A Critical Review of the Antimicrobial and Antibiofilm Activities of Green-Synthesized Plant-Based Metallic Nanoparticles.

Metallic nanoparticles (MNPs) produced by green synthesis using plant extracts have attracted huge interest in the scientific community due to their excellent antibacterial, antifungal and antibiofilm activities. To evaluate these pharmacological properties, several methods or protocols have been successfully developed and implemented. Although these protocols were mostly inspired by the guidelines from national and international regulatory bodies, they suffer from a glaring absence of standardization of the experimental conditions. This situation leads to a lack of reproducibility and comparability of data from different study settings. To minimize these problems, guidelines for the antimicrobial and antibiofilm evaluation of MNPs should be developed by specialists in the field. Being aware of the immensity of the workload and the efforts required to achieve this, we set out to undertake a meticulous literature review of different experimental protocols and laboratory conditions used for the antimicrobial and antibiofilm evaluation of MNPs that could be used as a basis for future guidelines. This review also brings together all the discrepancies resulting from the different experimental designs and emphasizes their impact on the biological activities as well as their interpretation. Finally, the paper proposes a general overview that requires extensive experimental investigations to set the stage for the future development of effective antimicrobial MNPs using green synthesis.

1. Introduction

Nanotechnology involves science, engineering and technology that considers matter at atomic, molecular or supramolecular levels to produce nanometric materials and nanosystems with improved properties, such as high surface-to-volume ratios and high dispersion in solution [1]. With sizes typically ranging between 1 and 100 nm in at least one dimension, nanomaterials and nanosystems can be synthesized by chemical, physical and/or biological methods [2,3].

In comparison to chemical and physical methods that can involve costly and toxic chemicals [2,4], the biological synthesis pathway, based on the use of biological sources (e.g., plants, bacteria, fungi and algae) represents an attractive option [5,6]. However, though these biological methods do not involve toxic chemicals in the preparation protocols, the microbial production of metal nanoparticles is highly demanding, time consuming and costly, requiring technology and practical microbiological experience to ensure cell culture and nanoparticle purification under aseptic conditions [7,8].

In contrast, the use of plants (e.g., extracts, fruit juices) for the synthesis of metal nanoparticles (MNPs) involves easy, simple, quick, environmentally friendly, sustainable and cost-effective processes, typically under moderate reaction conditions [9]. Plant-mediated synthesis of nanoparticles is also clinically adaptable and easily scalable for industrial production [10]. Interestingly, the secondary metabolites (e.g., polyphenols, flavonoids, tannins, terpenoids, alkaloids) contained in plant extracts often act as reducing and/or capping agents [11,12]. Depending on their morphological and physical characteristics (e.g., size, zeta potential), as well as their composition, MNPs from plants with medicinal value can exhibit improved antibacterial, antifungal and antibiofilm activities, thus constituting a very promising means of combating antimicrobial resistance [9,13,14,15,16].

Using different sources of metals (salts or oxide) and different plant extracts, the biological reduction method allows the synthesis of a large number of green MNPs, including silver (Ag), gold (Au), zinc oxide (ZnO), platinum (Pt), palladium (Pd), copper (Cu), iron oxide (Fe2O3 and Fe3O4), nickel oxide (NiO), magnesium oxide (MgO), titanium dioxide (TiO2) and indium oxide (In2O3) [8,14].

Investigation of plant systems as potential bio-factories for MNPs has received considerable attention, especially by researchers working in the field of phytonanotechnology, and pharmaceutical and clinical microbiology, as well as medicine [14,17]. Indeed, due to the surging popularity of green methods, more than 1000 research papers and reviews related to antibacterial, antifungal and antibiofilm properties of MNPs have been published to date (Pubmed and Google Scholar database). Most of the reviews published so far have mainly focused on predicting the antimicrobial mechanisms of MNPs and parameters that may influence their antibacterial, antifungal and activities, including: (i) the type (and origin) of plants used as bioreactor sources for biosynthesis, (ii) the reduction process of the metal salts (mainly silver, zinc and gold) used during the fabrication of nanoparticles, (iii) the particulate characteristics of MNPs (e.g., size, zeta potential and shape), as well as the characterization techniques allowing their determination, and (iv) the general protocols applied to evaluate the antimicrobial potential of metal-containing nanoparticles [10,13,14,17,18,19,20,21,22,23,24,25,26].

Unfortunately, it appears from these reviews that the methods used for assessing the antibacterial, antifungal and antibiofilm efficiency of MNPs are only partially elaborated in terms of standardization processes; therefore, it is hard to correlate or compare data from different studies to set out the product quality attributes and boost the development pipeline for high value antimicrobial nanoparticles. Hence, to provide an essential reference for readers, the present review briefly describes different in vitro and in vivo methods used for testing the antimicrobial activities of MNPs against planktonic bacteria, fungi and biofilms. The paper also presents tabular data (from 2010 to date) that summarize different in vitro experimental conditions used for assessing antimicrobial and antibiofilm activities of MNPs, including the major findings based on physicochemical characteristics (e.g., size and shape). We also discuss the discrepancies found in the obtained results. By doing so, this review guides scientists towards the most appropriate experimental settings for MNP evaluation and offers a useful resource for further complementary investigations. Moreover, it may pave the way for research and development of accessible and affordable drug formulations and wound-dressings containing green-synthesized metal and metal oxide nanoparticles.

2. Microbial Origins and Antimicrobial Resistance of Bacterial and Fungal Infections

The human body is not sterile; it is colonized by many microorganisms that are part of the normal microflora and live as harmless commensals [27]. Bacteria living under normal conditions on the skin, nasopharynx and intestine play an important protective role, as they prevent the growth of pathogenic microorganisms in these places. The loss of the protective functions of this barrier for any reason is an important factor in the onset of infections. As the body’s condition changes, immune bacteria that have previously been weakened can become pathogenic and cause infections ranging from minor to life-threatening [28].

Pathogenic microorganisms that are present in almost every location and environment on earth (e.g., air, soil biomass, water, plants and animals) can be sources of infections [29,30]. Contemporary lifestyles (e.g., imported food, air-conditioned environments, travel abroad and visits to hospitals) also contribute to the spread of infections. In other words, infections result from ever-changing interactions between microorganisms, the human as their host, and the environment around them [31].

Antimicrobial agents (antibiotics and antifungals) are used to treat or prevent different types of infections caused by bacteria and fungi (as well as certain parasites). However, most antimicrobials currently face the development and spread of antimicrobial resistance [32]. The scientific literature, and recent observations, show that a growing number of bacterial infections, such as tuberculosis, salmonellosis, pneumonia and gonorrhea are becoming more difficult to treat as the antibiotics used for their treatment become ineffective [33]. These situations lead to increased treatment failures, hospitalization time, economic burden and deaths [34]. The vast majority of antimicrobial drugs currently used promote genetic instability and increased mutagenesis in bacteria and fungi [35]. The spread and accumulation of antibiotic resistance bacteria (ARB) and antibiotic resistance genes (ARGs) in the environment are one of the greatest threats to human health and represent the emergence of a difficult situation in the world [36,37]. As antibiotic resistance is occurring naturally, ARG transmission also appears to be correlated with human activities [38]. ARB and ARGs have been found in various locations, such as ground- and drinking-water [39,40,41], agricultural soil, vegetables, sewage sludge, and agricultural products, such as fish [42,43,44,45]. Therefore, tackling this scourge is becoming a global emergency with an urgent need to overcome the ability of Gram-positive/negative bacteria and fungi to resist antimicrobial drugs [46]. Figure 1 summarizes the antibiotic microbial resistance attained by different micro-organisms through various intrinsic or acquired mechanisms [47].

Figure 1

Antibiotic resistance strategies in microorganisms. Mechanisms by which bacteria (and fungi) can resist antimicrobial molecules include: (i) target alterations and modifications through genetic mutations or post-translational modifications, (ii) increased active efflux of antibiotic out of the cell through efflux pumps, a type of membrane transporter located within the microbial membrane or wall, (iii) inactivation, destruction or degradation of the antibiotic through hydrolysis or modification by different enzymes (e.g., extended-spectrum β-lactamases) that can add specific chemical moieties, such as phosphoryl groups, (iv) decreased influx of antibiotic into the bacteria, e.g., through charges in the structure of the cell wall, (v) reduced permeability of the membrane that surrounds the bacterial cell. Created with BioRender.com (accessed on 1 March 2022).

It should be noted that misuse of antimicrobial drugs in both human and animal health may also represent an important source of antimicrobial resistance. Antimicrobial agents are extensively used in animal husbandry and agriculture for prophylactic and therapeutic purposes [48]. In addition, antimicrobial drugs are used as growth promoters in animal feed, and to increase crop productivity. Such overuse of antimicrobial drugs outside of clinics has contributed tremendously to the rise in antimicrobial resistant strains and led to the growing need for new antimicrobial agents that can effectively treat and prevent infectious diseases worldwide. Therefore, implementing the One Health approach is proving to be a great necessity, even of the utmost urgency [49,50].

In nature, microorganisms rarely live in isolated colonies of the same species. They are characterized by “life” in biocenoses called biofilms [51]. Biofilms are communities of microbes attached to surfaces, which can be found in medical, industrial and natural settings. In fact, life in a biofilm probably represents the predominant mode of growth for microbes in most environments [52,53].

Mature biofilms have a number of distinct characteristics. They are surrounded by an extracellular matrix that contains polysaccharides, nucleic acids, lipids, proteins, water and ions [53,54]. This matrix provides structure and protection to the community of microorganisms. Microbes growing in a biofilm also have a characteristic architecture, generally comprised of macrocolonies (containing thousands of cells) surrounded by fluid-filled channels. Biofilm-grown microbes are notorious for their resistance to various antimicrobial agents, including clinically relevant antibiotics [55,56,57].

Biofilms are formed in five stages (Figure 2; [58]). The first stage (attachment) is related to the primary adhesion and adsorption of bacteria or fungi; this step is a reversible process (stage 1). In the second stage (so-called fixation or colonization), microorganisms secrete polymers that ensure strong and irreversible adhesion (stage 2). In the proliferation stage, the microorganisms attach to the epithelium, facilitate the attachment of new microbial cells, and connect the entire colony to the intercellular filler. With the accumulation of nutrients, microorganisms begin to multiply (stage 3). The following step is the stage of full maturation; the formed biofilms acquire their size and shape, and the intercellular filler protects them from external negative factors (e.g., oxygen, temperature, nutrient) (stage 4). Finally, in the last stage, bacterial spread (dispersion) occurs, during which individual bacterial cells periodically separate, creating new colonies (stage 5) [58,59,60,61]. All surfaces, including medical equipment, irrespective of their nature and the environment in which they are used, are susceptible to the colonization and infection of microorganisms and the formation of biofilms [56].

Figure 2

Developmental stages involved in microbial biofilm formation (From [58] with permission from Frontiers in Microbiology).

Overall, biofilms do not only represent a bacterial layer of mucus, but also a biological system composed of bacteria and/or fungi that are organized into a coordinated and functional biocenosis through intercellular chemical communication or quorum sensing [61]. Microorganisms in biofilms form metabolic consortia that are characterized by (i) food exchange between microbial cells (i.e., each microorganism becomes a food source for another), (ii) primitive exchange of genetic information, and (iii) resistance to phagocytosis and antibiotics. Therefore, biofilm-forming bacteria (or fungi) can be 100-fold more resistant to antimicrobial agents and disinfectants than planktonic bacteria [59]. Recurrent biofilm-induced infections are especially dangerous to people with health problems, as they can cause hospital infections. In USA, one out of thirty-one hospitalized patients have at least one healthcare-associated infection [62]. In Europe, approximately 5% of hospitalized patients suffer from hospital infections, i.e., 4.1 million people every year, including about 37 thousand deaths [63,64,65]. The prevalence of healthcare-associated infections can reach 20% in middle- and low-income countries. In addition, 4 to 56% of hospital-born babies die from health care-associated infections in the neonatal period in developing countries. This prevalence can reach 75% in south-east Asia and sub-Saharan Africa [63].

3. General Background on Antimicrobial and Anti-Biofilm Metallic Nanoparticles Green-Synthesized

3.1. Green Synthesis of MNPs

In general, there are two ways of synthesizing MNPs: the “bottom-up” and “top-down” approaches. Both methods of synthesis can be performed either in liquid (e.g., water, ethanol, hexane, toluene, ethylene glycol and others), gas, solid, or supercritical fluids, or in a vacuum [17,66,67].

In bottom-up synthesis, atoms, molecules or clusters are grouped to form nanostructured materials [3,17,68]. A wide variety of physical and chemical methods belong to this category. The physical methods include spinning, physical vapor deposition and molecular-beam epitaxy (Figure 3) [7,69]. The chemical methods comprise sol-gel processes, laser pyrolysis, chemical vapor deposition, aerosol-based processes, atomic or molecular condensation and precipitation, plasma-spraying synthesis and supercritical fluid technology [14,66,67,70]. The green synthesis of MNPs is also a class of bottom-up methods in which reduction and oxidation are the foremost chemical reactions [2,21]. Conversely, in the top-down approach, the bulk materials are broken down gradually (or collapsed little-by-little) to yield nanoparticles. Mechanical milling, sputtering and lithography belong to physical top-down methods, while chemical top-down methods include electro-explosion and chemical etching [14,17,66,68].

Figure 3

Overview of the synthesis of metal nanoparticles.

In general, chemical and physical methods have the reputation of being expensive and poorly accessible to all scientists across the world, especially in developing countries [2,3]. The chemical methods are also harmful to the environment due to the use of organic solvents and highly reactive toxic chemicals and reducing agents [71]. The latter can generate unwanted by-products that can, in turn, cause potential environmental and biological risks [39,40,67,72,73]. The physical methods are limited by low production rates and high energy consumption, as well as requiring sophisticated equipment and stringent conditions [14].

On the other hand, the synthesis pathways using plant extracts or microorganisms are environmentally benign, non-toxic, cost-effective, simple and easily up-scalable for industrial production [23,67,74,75,76,77,78]. Using renewable materials and mild solvent media, biogenic methods of synthesis of MNPs offer additional benefits, including one-pot synthesis, no need for catalyst use, clean, yet straightforward, reaction conditions, and no toxic waste generation [14,21]. Nevertheless, synthetic microbial methods include several disadvantages, such as microorganism cultivation and the optimization of different growth parameters (e.g., nutrient medium, salt concentration, temperature, pH, incubation time, inoculum quantity, etc.) [79]. A combination of different physical factors, such as light, ultrasonic waves, microwaves, heating, etc., is also required to produce MNPs by synthetic microbial methods [36,42,80,81]. As a result, microbe-mediated synthesis of MNPs requires expertise and is also time-consuming [17]. Moreover, the intrinsic ability of microorganisms to act as reducing and capping agents and contribute to the amalgamation of metal ions into MNPs is lower than that of plant metabolites [82].

MNPs synthesized by chemical (or physical) synthesis can exhibit antibacterial activity, as described previously [83,84,85,86]. However, many researchers have reported that green-produced MNPs showed higher activity than those produced chemically [4,87]. This is in most cases the result of a kind of synergy between the intrinsic activity of the metals and of the plant metabolites themselves.

3.1.1. Plant-Mediated Synthesis of Silver, Gold and Zinc Oxide Nanoparticles

The green synthesis of silver nanoparticles (AgNPs) is typically achieved by combining plant extracts and silver halides, such as silver bromide (AgBr), silver chloride (AgCl), and silver iodide (AgI) [88,89].

During the biosynthesis of green ZnO nanoparticles, zinc nitrate hexahydrate (Zn(NO3)2·6H2O) or zinc acetate (Zn(OOCCH3)2) are typically used as precursors while different plant extracts act as bio-reductants [90,91].

To synthesize gold nanoparticles (AuNPs) using green sources, plant extracts are simply mixed with solutions of gold salts, such as auric chloride (AuCl3) and chloroauric acid (HAuCl4) [92,93].

3.1.2. Synthesis of Platinum and Palladium Nanoparticles Using Plant Extracts

Bio-fabricated PtNPs can be obtained using plant extracts as eco-friendly reducing reagents, along with sodium tetrachloroplatinate (II) (Na2PtCl4) and chloroplatinic acid hexahydrate (H2PtCl6·6H2O) as precursor salts [90,91]. In the process of the bio-fabrication of PdNPs, plant extracts are mixed with palladium chloride (PdCl2) or palladium acetate (Pd(OAc)2). Depending on their intrinsic properties, the phytocomponents from the plant extracts reduce the Pd ions into atoms and then produce metallic Pd nanoparticles [90,91,94].

3.1.3. Biosynthesis of Other Green Metallic Nanoparticles

Nanosized CuO particles can be obtained by reducing cupric salts (e.g., cupric chloride (CuCl2), cupric sulfate (CuSO4) and cupric nitrate (Cu(NO3)2) using phytoconstituents from plant extracts [90,91].

The precursor salts for the green synthesis of Fe2O3 (hematite) and Fe3O4 (magnetite) nanoparticles include ferric chloride hexahydrate (FeCl3·6H2O), ferric nitrate nonahydrate (Fe(NO3)3·9H2O), ferric acetylacetonate (Fe(C5H8O2)3), and ferrous sulfate (FeSO4) [95,96]. Nickel nitrate hexahydrate (Ni(NO3)2·6H2O) and magnesium nitrate hexahydrate (Mg(NO3)2·6H2O) can be used as precursors for the synthesis of NiO and MgO nanoparticles by green process, respectively [90,91,97]. Additionally, nickel chloride (NiCl2) and indium nitrate (In(NO3)3·H2O) can be reduced to form nickel oxide nanoparticles and indium oxide (In2O3), respectively, by plant extracts [98,99]. Lead nanoparticles can be synthesized by green methods using lead oxalate (Pb(COOH)2) or lead nitrate (Pb(NO3)2) as metal precursors [100,101].

Concerning TiO2 NPs, their green synthesis can be successfully achieved by mixing plant extracts as reducing/capping materials with metatitanic acid (TiO(OH)2) or titanium tetraisopropoxide (Ti[OCH(CH3)2]4) as precursors [90,91]. Additionally, numerous bimetallic and trimetallic nanoparticles are currently synthesized by green methods. This is the case for Ag/Pt NPs, Pd/Fe3O4 NPs, Fe3O4/MgO NPs, ZnO/CoO NPs, ZnO/MnO NPs, Au/Pt/Ag NPs and Cu/Cr/Ni NPs obtained by mixture of the corresponding precursors and plant extracts [102,103,104,105].

Over the last decade, other antimicrobial hybrid derivatives, such as nanocellulose/metal and nanocellulose/oxide metal, have attracted the attention of several researchers and industries due to their environmentally friendly status and low cost compared to synthetic polymer- (e.g., polylactic-co-glycolic acid (PLGA), polyvinylalcohol (PVA), poly(ethylene glycol) methyl ether-block-poly(lactide-co-glycolide) (mPEG-PLGA), chitosan, gelatin) encapsulated MNPs [106,107,108,109,110,111,112,113]. Cellulose is a ubiquitous natural polymer which can be produced from a broad range of biomass. It is the key component in natural fibers and an excellent candidate for synthesis of bio-based materials due to its various physicochemical properties, including biodegradability, biocompatibility, environmental friendliness, renewability, affordability and colloidal stability [114,115].

Nanocellulose is defined as cellulose material that has been broken down into particles of less than 100 nm [116]. It is essentially produced through chemical or mechanical action on the plant cellulose or bacterial cellulose. Nanocellulose is classified into cellulose nanocrystals, cellulose nanofibers and bacterial nanocellulose [108,114,115]. Apart from plant materials (e.g., wood, oil palm biomass, bamboo, rice husk, coconut husk), cellulose nanocrystals and cellulose nanofibrils can also be extracted from tunicate, a type of marine invertebrate. In contrast, bacterial nanocellulose is a growing nanoproduct that can be obtained through several kinds of mutual fermentation bacteria, such as Gluconacetobacter xylinus [115,117].

Nanocellulose does not inherently elicit any antimicrobial activity. However, by functionalization with green metal/metal oxide nanoparticles (e.g., Au, Ag, Cu, CuO, MgO, ZnO, Fe3O4 and TiO2), it is possible to endow nanocellulose composites with antimicrobial properties (Figure 4) [118,119]. These biosynthesized metal-based antimicrobial agents may also exhibit good efficacy and resilience towards microbial resistance [114,117]. In this context, Mocanu et al. [120] recently demonstrated the impact of functionalization of bacterial nanocellulose with ZnO NPs, green-synthesized using propolis extract, on antimicrobial activity against Bacillus subtilis and Candida albicans, in comparison to ZnO NPs and the extract alone. Additionally, Razavi et al. [121] prepared antimicrobial bacterial nanocellulose film decorated with silver, copper and palladium MNPs biosynthesized using mulberry fruit (Morus alba L.) extract. Due to their significant activity against Escherichia coli and Listeria monocytogenes, the authors suggested that the fabricated nanocomposite film might be useful as a novel biomedical treatment to combat pathogens on food commodities.

Figure 4

Illustrative presentation of green synthesis of nanocellulose/metal or metal oxide hybrid nanocomposites. (I) Different types of nanocellulose in dispersion; (II) Electron microscope images of MNPs in cellulose (a and b for SEM images, and c and d for TEM images, respectively): white and black arrows point to MNPs and defibrillated cellulose, respectively (Adapted from [107,118] with permission from The Royal Society of Chemistry and Scientific Reports).

Green-synthesized plant-based MNPs can be characterized using a wide range of physicochemical tools, such as UV-visible absorption spectroscopy (for metal surface plasmon resonance), zeta potential (for surface-charge determination), transmission electron microscopy (for size and shape analysis), X-ray diffraction (for crystallinity assessment), energy-dispersive X-ray spectroscopy (EDX) (for the determination of elemental composition on the surface), Fourier transform infrared spectroscopy (FTIR) (for the detection of organic functional groups of phytoconstituents), thermogravimetric analysis (for thermal stability), and Raman spectroscopy (for surface-capping tracking) [9,68,122,123,124].

3.2. Antimicrobial Activities of Green-Synthesized MNPs

Plant-mediated synthesis imparts several advantages to MNP technology for the development of alternative products against infectious diseases. Indeed, most of the green MNPs from plant-derived materials are highly effective and non-specific antimicrobial agents, showing remarkable activity against the growth of a broad spectrum of bacterial and fungal species, in both planktonic and biofilm forms, including nosocomial and multi-drug-resistant strains (Tables 1–8 and S1–S7) [11,13,14,16,125].

Given their nano-particulate features, biosynthesized MNPs provide a large surface area that increases their interactions with microorganisms, thereby resulting in strong antimicrobial activity. The antimicrobial properties of green MNPs also depend on their particle shape. Moreover, the variety of green reagents (plant extracts or phytoconstituents), metal precursors and synthetic conditions (e.g., physicochemical parameters) used have a significant effect on the antimicrobial activity of MNPs [12,13,126,127].

Biofabricated MNPs may act in different ways, including: (i) destruction of the microbial cell wall, (ii) damage to efflux pump mechanisms, (iii) inhibition of deoxyribonucleic acid (DNA) replication and enzyme functions, (iv) ribosome disintegration, (v) generation of reactive oxygen species (ROS) and induction of oxidative stress; (vi) triggering of both innate and adaptive host immune responses, and (vii) inhibition of biofilm formation (Figure 5) [47,128,129]. The mechanisms of action of MNPs depend on their origin as well as their biological, physical and chemical properties [130].

Figure 5

Different mechanisms of action of MNPs in microbial cells. The combination in a single nanomaterial of a multitude of cellular effects may have a tremendous impact in fighting multi-drug-resistant microorganisms (From [129] with permission from Frontiers in Microbiology).

The antibacterial, antifungal and antibiofilm activities of the most biosynthesized MNPs are briefly summarized in the following paragraphs. In addition, the unique physicochemical characteristics of these MNPs are briefly described since their antimicrobial activities are also attributed to their size, high surface area, zeta potential and shape. For more details, the reader can refer to more specific reviews [10,13,14,17,23,24,25,26,128,131,132,133,134].

3.2.1. Silver Nanoparticles

Elemental silver has been widely used as an antimicrobial agent since ancient times [89]. To improve their antibacterial activity and reduce their toxicity, silver ions can be transformed into metallic silver nanoparticles through biological and biomimetic methods of synthesis [9].

Details of the antimicrobial and anti-biofouling activities of these bioactive nanoparticles are given in Table 1 and Table 2. It is notable that green AgNPs have demonstrated the ability to reduce microbial infections in skin and burn wounds and to prevent bacterial colonization on the surface of various medical devices, such as catheters and prostheses. Acting as capping agents, different multi-functional phytochemicals contribute efficiently to these antimicrobial activities [74,78,82,128]. Moreover, AgNPs can operate synergistically with standard antibiotics, such as gentamycin and streptomycin [129,135,136,137]. Hence, these combinations can be used effectively against antibiotic-resistant pathogens. Additionally, the antifungal activity of AgNPs has been extensively studied and demonstrated in the literature [138,139].

Table 1

Green silver nanoparticles exhibiting antimicrobial activity.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics(Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density,Positive Control	Tested Bacteria and Fungi	MIC, ZOI or PI *
Lysiloma acapulcensis	Roots	Silver nitrate 1 mM/plant extract 2% (1:1 v/v)Room temperature 2 min pH NM	Spherical 1.2–62 nm	Diffusion 37 °C24 hpH NM Inoculum NMNo control	E. coli ATCC 25922P. aeruginosa ATCC 27853S. aureus ATCC 49476	18 15 16 mm	[87]
Perilla frutescens	Leaves	Silver nitrate 2 mM/plant extract 10% (9:1 v/v) 50 °C2 h pH NM	Spherical, rhombic, triangle, and rod25.7 nm	Diffusion 37 °C24 hpH NMInoculum NMStreptomycin **	E. coli B. substilis S. aureus	1412 10 mm	[149]
Ocimum canum	Leaves	Silver nitrate 1 mM/plant extract 10% (9:1 v/v)80 °C15 min pH NM	Spherical 6.1–32.1 nm	Diffusion 28 °C24 hpH NMInoculum NMNo control	E. coli	25 mm	[150]
Piper longum	Catkin	Silver nitrate 1 mM/plant extract 10% (5:1 v/v)Room temperature 2 h pH NM	Spherical 10–42 nm	Diffusion 37 °C24 hpH NMInoculum NMNo control	B. cereus MTCC 1272E. coli MTCC 1687K. pneumoniae MTCC 530Proteus mirabilis MTCC 425P. aeruginosa MTCC 1688S. typhi MTCC 531S. aureus MTCC 96	12 13 14 15 1112 11 mm	[139]
Diospyros malabarica	Fruits	Silver nitrate 1 mM/plant extract 20% (9:1 v/v)25 °C 1 hpH NM	Spherical 17.4 nm	Diffusion 37 °C24 hpH NMInoculum NM Streptomycin 10 µgTetracycline 30 µgChloramphenicol 30 µg	E. coli S. aureus	13 12 mm	[151]
Pyrenacantha grandiflora	Tuber	Silver nitrate 1 mM/plant extract 0.1% (1:1 v/v)Room temperature Incubation time NMpH NM	Spherical3–25 nm	Dilution 37 °C24 h pH NMInoculum NMNo control	E. coli K. pneumoniae S. aureus	0.8 0.8 0.8 µg/mL	[152]
Carissa carandas	Leaves	Silver nitrate 1 mM/plant extract 10% (9:1 v/v)60 °C1 h pH 7.2	Spherical 30 nm	Diffusion 37 °C24 hpH NM1 × 108 CFU/mL No control	S. typhiEnterococcus faecalisShigella flexneriCitrobacter spp.Gonococci spp.	12 16 24 14 21 mm	[153]
Solanum tricobatum	Leaves	Silver nitrate 1 mM/plant extract 1.5% (1:10 v/v) 37 °C24–48 h pH NM	Irregular 26.5 nm	Diffusion 35 °C18 h pH NMInoculum NMNo control	S. aureus P. aeruginosa E. coli K. pneumoniae	3012 14 18 mm	[154]
Melissa officinalis	Leaves	Silver nitrate 5 mM/plant extract 25% (1:2 v/v)25 °C1 hpH NM	Spherical12 nm	Diffusion36 °C24 hpH NMInoculum NMNo control	E. coli S. aureus	1213 mm	[155]
Piper betle	Leaves	Silver nitrate 1 mM/plant extract 10% (10:1 v/v)Room temperature24 hpH NM	SphericalSize NM	Diffusion 30 °C24 hpH NMInoculum NMNo control	B. subtilis Klebsiella planticola	14 13 mm	[156]
Rosa canina	Fruits	Silver nitrate 1 mM/plant extract% NM (5:1 v/v)85 °CIncubation time NMpH NM	Spherical 13–21 nm	Dilution 37 °C24 h pH NM2.4 × 107 CFU/mLNo control	Bacillus cereusE. coli ATCC 10536S. aureus ATCC 6538P. aeruginosa ATCC 9027Enterococcus hirae ATCC 10541Legionella pneumophila ATCC 33152	32 256 256 128 256 16 µg/mL	[157]
				Dilution 25 °C48 h pH NM2.4 × 107 CFU/mLNo control	C. albicans	128 μg/mL
Fagonia indica	Callus	Silver nitrate 4 mM/plant extract 2% (1:1 v/v)20 °C3 hpH NM	Cubic Size NM	Diffusion 37 °C24 hpH NM1 × 108 CFU/mLCiprofloxacin **	E. coli ATCC 23716S. typhi ATCC 35664Shigella sonnei ATCC 29930Citrobacteramalonaticus ATCC 25405	12 13 1312 mm	[158]
Barleria longiflora	Leaves	Silver nitrate 1 mM/plant extract 20% (9:1 v/v) Temperature NMIncubation time NMpH NM	Spherical 2.4 ± 0.5 nm	Diffusion 37 °C24 hpH NMInoculum NM Chloramphenicol **	Enterococcus spp.Streptococcus spp.Bacillus megateriumPseudomonas putidaP. aeruginosaS. aureus	18 161517 18 14.5 mm	[159]
Ipomoea batatas	Outer peels	Silver nitrate 1 mM/plant extract 40% (10:1 v/v)55 °C24 hpH NM	Shape NMSize NM	Diffusion Temperature NMIncubation time NMpH NMInoculum NMNo control	Enterococcus feacium DB 01 S. enteritica KCCM 11806Listeria monocytogenes ATCC 19111B. cereus KCTC 3624S. aureus ATCC 13565	1011 11 11 0 mm	[160]
Oedera genistifolia	Leaves	Silver nitrate 0.1 mM/plant extract 20% (9:1 v/v) Room temperature1 hpH NM	Spherical 10–60 nm	Dilution 37 °C 24 h pH NM1 × 108 CFU/mLCiprofloxacin **	Enterobacter cloacae ATCC 13047Listeria ivanovic ATCC 19119Streptococcus uberis ATCC 700407S. aureus ATCC 29213Vibrio spp.Mycobacterium smergatis ATCC 19420	0.5 10.50.5 0.250.25 mg/mL	[161]
Derris trifoliate	Seeds	Silver nitrate 1 mM/plant extract 20% (20:1 v/v)Temperature NMIncubation time NMpH NM	Spherical 16 ± 5 nm	Diffusion NM24 hpH NMInoculum NMNo control	E. coli MTCC 723 K. pneumoniae MTCC 109 P. aeruginosa MTCC 424S. aureus MTCC 96	19.520360 mm	[162]
Ficus krishnae	Stem bark	Silver nitrate 1 mM/plant extract 5% (1:1 v/v)37 °C24 h pH NM	Spherical 160–260 nm	Diffusion 37 °C24 h pH NMInoculum NMNo control	E. coli MTCC 45S. typhimurium MTCC 98S. aureus ATCC 29122	18 13 12 mm	[163]
Psidium guajava	Leaves	Silver nitrate 10 mM/plant extract 2% (10:1 v/v)70 °C1 h pH NM	Spherical 96 ± 4 nm	Diffusion 37 ± 2 °C48 hpH NM 1–2 × 105 CFU/mLNo control	C. albicans ATCC 10231	14.2 mm	[164]
Citrus limon	Leaves	Silver nitrate 2 mM/plant extract 20% (9:1 v/v) 25 °C1 hpH NM	Spherical8–15 nm	Diffusion Temperature NM 18–24 hpH NMInoculum size NMNo control	Fusarium oxysporium Alternaria brassicicola	15 10 mm	[165]
Chaenomeles sinensis	Fruits	Silver nitrate 1 mM/plant extract 10% (ratio NM)80 °C65 min pH NM	Spherical 5–20 nm	Diffusion37 °C24 h pH NMInoculum NM Neomycin **	E. coli S. aureus	1410 mm	[166]
Persicaria odorata	Leaves	Silver nitrate 1 mM/plant extract 2% (10:1 v/v)25 °C24 hpH NM	Spherical 11 ± 3 nm	Dilution 37 °C18 h pH NM1 × 106 CFU/mLNo control	S. epidermidis ATCC 12228MRSA ATCC 43300	3-LR ***6-LR	[167]
Citrus reticulata	Peels	Silver nitrate 1 mM/plant extract 21.8% (1:1 v/v)Temperature NMIncubation time NMpH NM	Spherical45 nm	Dilution 37 °C48 hpH NM1 × 105 CFU/mLNo control	Desulfovibrio spp.	3-LR	[168]
Cuccuma longa	Rhizome	Silver nitrate 1 mM/plant extract 6.8% (4:1 v/v)Temperature NM24 hpH NM	Spherical 18 nm	Dilution 37 °C24 h pH NM1 × 108–109 CFU/mLNo control	E. coli Listeria monocytogenes	4-LR4-LR	[169]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned. *** LR = log reduction. A 1-log, 2-log, 3-log, 4-log, 5-log and 6-log reduction in living microorganisms or CFUs by MNPs corresponds to their inactivation or inhibition of 90, 99, 99.9, 99.99, 99.999 and 99.9999%, respectively. NM = not mentioned.

Table 2

Green silver nanoparticles with antibiofilm activity.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics(Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density,Positive Control	Tested Bacteria and Fungi	MIC, ZOI or PI *
Punica granatum	Peel	Silver nitrate */plant extract 5% (ratioNM)Temperature NMIncubation time NMpH NM	Spherical 32–85 nm	Microtiter plate 37 °C24 h pH NM1.5 × 108 CFU/mLNo control	P. aeruginosa ATCC 10662	89.6%	[170]
Artemisia scoporia	NM	Silver nitrate 1000 mM/plant extract 10% (20:1 v/v) Temperature NM24 h pH NM	Spherical 10–80 nm	Microtiter plate 37 °C24 h pH NM1.5 × 108 CFU/mLNo control	S. aureus	6.25 µg/mL	[171]
Prosopis juliflora	Leaves	Silver nitrate 1 mM/plant extract 10% (9.5:0.5 v/v) 25 °C40 min pH NM	Spherical10–20 nm	Congo red agar plate37 °C24–48 hpH NMInoculum size NMNo control	B. substilis P. aeruginosa	NMNM	[172]
Malva sylvestris	Leaves	Silver nitrate 1 mM/plant extract 20% (10:0.4 v/v) Temperature NMIncubation time NMpH NM	Spherical 10–50 nm	Dilution 37 °C40 h pH NM1 × 108 CFU/mLNo control	P. aeruginosa 48P. aeruginosa B 52	62.562.5 μg/mL	[173]
Cannabis sativa	Stem	Silver nitrate 1 mM/plant extract 10% (1:1 v/v) Temperature NMIncubation time NMpH NM	Spherical 20–40 nm	Microtriter plate 37 °C24 hpH NM 2–5 × 106 CFU/mL No control	P. aeruginosa PA01E. coli UTI89S. epidermidis	6.2512.550 µg/mL	[174]
Rhodiola rosea	Rhizome	Silver nitrate 5 mM/plant extract 10% (2:8 v/v) 90 °C10 min pH NM	Spherical 15–30 nm	Dilution 37 °C 24 hpH NM 1–2 × 106 CFU/mLNo control	P. aeruginosa E. coli	50100 µg/mL	[131]
Flacourtia indica	Leaves	Silver nitrate 1 mM/plant extract 10% (1:1 v/v) 70 °CIncubation time NMpH NM	Spherical 45.9–64.9 nm	Congo red 37 °C24 hpH NMInoculum size NMNo control	Acinetobacter baumannii SAB5P. aeruginosa ETPS11K. pneumoniae SKP7P. mirabilis PPM8E. coli ETEC12	80 80808080 μg/mL	[116]
Dodonaea viscosa	Leaves	Silver nitrate 1 mM/plant extract 10% (ratio NM)Temperature NM18 hpH NM	Spherical 40–55 nm	Crystal violet assay 37 °C24 hpH NM1 × 107 CFU/mLNo control	C. albicans Candida tropicalis Candida glabrata	808080%	[175]
Piper betle	Leaves	Silver nitrate 1 mM/plant extract 5% (19:1 v/v) 37 °C6 hpH NM	Spherical 156.4 nm	Microtiter plate18 °CTemperature NMpH NMInoculum size NMNo control	Serratia marcescens Proteus mirabilis	7169%	[176]
Pedalium murex	Seed	Silver nitrate 1 mM/plant extract 5% (49:1 v/v) Temperature NM20 min pH NM	Hexagonal20–30 nm	Microtiter plate 37 °C24 hpH NMInoculum size NMNo control	Enterococcus faecalis S. aureus Shigella sonnei P. aeruginosa	6462 5054%	[177]
Solanum nigrum	Fruit	Silver nitrate 1 mM/plant extract 10% (50:1 v/v) Temperature NM10 min pH NM	Spherical 10–20 nm	Microtiter plate 37 °C24 hpH 7.2 1 × 108 CFU/mLNo control	Bacillus pumulis Enterococcus faecalis Proteus vulgaris Vibrio parahaemolyticus	92847462%	[178]
Eucalyptus globulus	Leaves	Silver nitrate 1 mM/plant extract 20% (4:1 v/v) 60 °C 30 min pH 8	Spherical 18 nm	Microtitre plate37 °C24 hpH NM1 × 107 CFU/mLNo control	P. aeruginosa S. aureus	95 90%	[179]
Allophylus cobbe	Leaves	Silver nitrate 5 mM/plant extract 20% (10:1 v/v) 60 °C6 hpH 8	Spherical 2–10 nm	Microtitre plate 37 °C4 h pH NM1 × 106 CFU/mL No control	P. aeruginosa Shigella flexneri S. aureus Streptococcus pneumoniae	90906075%	[180]
Cinnamomum aromaticum	NM	NMNMNMNM	Spherical 15–50 nm	Dilution Temperature NM24 hpH NM 1 × 106 CFU/mL No control	Streptococcus agalactiae ATCC 27956	4 μg/mL	[181]
Prunica granatum	Leaves	Silver nitrate 1.5 mM/plant extract 5% (1:1 v/v)31.4 °C20 min pH NM	Spherical 37.5 nm	Congo red agar 37 °C 24 hpH NM Inoculum size NMNo control	P. aeruginosa S. aureus	4528%	[182]
Terminalia catappa	Leaves	Silver nitrate 10 mM/plant extract 5% (1:1 v/v)30 °C20 min pH NM	Spherical 3.5–10.1 nm	Microtiter plate 37 °C24 h pH NM 1 × 107 CFU/mL No control	P. aeruginosa S. aureus	73.769.6%	[183]
				Microtiter plate 37 °C24 h pH NM 5 × 106 CFU/mL No control	C. albicans	63.6%

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. The quantity or concentration is not mentioned. NM = not mentioned.

In the context of the fight against antibiotic resistance, green-synthesized AgNPs may be used as vehicles to transport oligonucleotide-based antimicrobials [140,141,142]. AgNPs can also be incorporated in hydrogel beds, cyclodextrins, and lipid-based formulations (e.g., liposomes), creating the potential for controlled release and targeted delivery [143,144,145]. Interestingly, AgNPs are found in a number of commercially available products, including medical devices for healthcare settings, dietary and health supplements, potential additives to animal feed, food packing materials and kitchen appliances [146,147,148].

3.2.2. Gold Nanoparticles

Due to their outstanding antimicrobial and antibiofilm properties, biosynthesized AuNPs are considered one of the most attractive MNPs. Indeed, as shown in Table 3, green AuNPs significantly inhibit the growth of medically important pathogenic bacteria and fungi.

Table 3

Green gold nanoparticles exhibiting antibacterial and antifungal activities.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics(Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density,Positive Control	Tested Bacteria and Fungi	MIC, DOI or PI *
Piper betle	Leaves	Gold (III) chloride 1 mM/plant extract 1% (10:1 v/v)30 °C 24 h pH NM	Spherical Size NM	Diffusion30 °C24 hpH NMInoculum size NMNo control	B. subtilis Klebsiella planticola	1314 mm	[156]
Musa acuminata	Flowers	Chloroauric acid 1 mM/plant extract 25% (9:1 v/v)Room temperature30 min pH NM	Spherical10–16 nm	Diffusion Temperature NM24 hpH NMInoculum size NMStreptomycin 10 μg	S. aureus Enterococcus faecalis E. coli S. typhi P. aeruginosa Proteus mirabilis	0117998 mm	[192]
Zingiber officinale	Roots	Chloroauric acid 1 mM/plant extract 1% (2:1 v/v)50 °C 24 hpH NM	Hexagonal10–20 nm	Diffusion37 °C 24 hpH NM1.5 × 108 CFU/mLNo control	S. aureus E. coli K. pneumoniae	141117 mm	[193]
Areca catechu	Nut	Chloroauric acid 1 mM/plant extract 5% (10:1 v/v)80 °C1 hpH NM	Spherical 14 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	S. aureus E. coli	1214 mm	[194]
Momordica cochinchinensis	Rhizome	Chloroauric acid 0.01 mM/plant extract 10% (2:1 v/v)Room temperature24 hpH NM	Spherical 16 ± 2 nm	Diffusion37 ± 1 °C24 hpH NM1 × 108 CFU/mLStreptomycin 100 µg/mL	S. aureus E. coli B. subtilis P. aeruginosa	19221924 mm	[195]
Plumeria alba	Flowers	Chloroauric acid 1 mM/plant extract 5% (5:2 v/v)Room temperature4 hpH NM	Spherical 15 nm	Diffusion37 °C 24 hpH NMInoculum size NMNo control	E. coli	16 mm	[196]
Coleus forskohlii	Root	Chloroauric acid 0.1 mM/plant extract 8% (1:1 v/v)Room temperature2 hpH 13	Spherical 5 nm	Diffusion37 °C 24–48 hpH NMInoculum size NMTetracyclin 30 μg/mL	Proteus vulgaris Micrococcus luteus	1814 mm	[197]
Euphorbia wallichii	Leaves	Chloroauric acid 1 mM/plant extract 5% (1:10 v/v)30 °C24 hpH NM	Hexagonal8 nm	Dilution34 °C 24 hpH NMInoculum size NMStreptomycin **	E. coli S. aureus Bacillus pumilus P. aeruginosa K. pneumonia	2115211717 mm	[198]
Coleus aromaticus	Leaves	Chloroauric acid 1 mM/plant extract 30% (1:1 v/v)100 °C30 minpH NM	Triangular20 nm	Diffusion37 °C 24 hpH NM1 × 108 CFU/mLNo control	S. epidermidis E. coli	2227 mm	[199]
Origanum vulgare	Leaves	Chloroauric acid 1 mM/plant extract 10% (10:1 v/v)85 °C1 min pH NM	Spherical52 nm	Diffusion37 °C 24 hpH NM1 × 108 CFU/mLNo control	Salmonella enteritidis ATCC 13076E. coli ATCC 25922Listeria monocytogenes ATCC 13932S. aureus ATCC 6538C. albicans ATCC 10231	108102128 mm	[200]
Perilla frutescens	Leaves	Chloroauric acid 1 mM/plant extract 10% (1:10 v/v)30 °C10 minpH NM	Triangular50 nm	Diffusion37 °C 24 hpH NMInoculum size NMNo control	E. coli B. subtilis S. aureus	141010 mm	[201]
Parkia roxburghii	Leaves	Chloroauric acid 1 mM/plant extract 1% (1:1 v/v)30 °C12 hpH NM	Quasi-spherical 5–25 nm	Diffusion37 °C 24 hpH NMInoculum size NMNo control	S. aureus E. coli	NMNM	[202]
Cibotium barometz	Roots	Chloroauric acid 1 mM/plant extract 5% (20:1 v/v)80 °CIncubation time NMpH NM	Spherical 23 nm	Diffusion37 °C 24 hpH NMInoculum size NMNeomycin 30 µg	E. coli ATCC 10798S. aureus ATCC 6538Salmonella enterica ATCC 13076P. aeruginosa ATCC 10145	16171312 mm	[203]
Mangifera indica	Seed	Chloroauric acid 1 mM/plant extract 10% (6:4 v/v)80 °C1 hpH NM	Spherical50 nm	Diffusion37 °C 24–48 hpH NM1 × 108 CFU/mLNo control	E. coli S. aureus	2525 μg/mL	[204]
Rhodiola rosea	Rhizome	Chloroauric acid 1 mM/plant extract 10% (10:1 v/v)80 °C30 minpH NM	Spherical 13–17 nm	Diffusion37 °C 24 hpH NMInoculum size NMNo control	S. aureus ATCC 29213E. coli ATCC 25922	15 12 mm	[131]
Amomum villosum	Fruit	Chloroauric acid 1 mM/plant extract 10% (10:1 v/v)100 °C 60 minpH NM	Spherical 5–10 nm	Diffusion37 °C 24 hpH NMInoculum size NMNeomycin **	S. aureus E. coli	1515 mm	[205]
Syzygium cumini	Seed	Chloroauric acid 1 mM/plant extract 2% (1:2 v/v)90 °C 1 hpH NM	Spherical 13–30 nm	Diffusion32 °C 24 hpH NM1 × 104 CFU/mLGentamicin **	E. coli B. subtilis S. aureus	303329 mm	[206]
Hovenia dulcis	Fruit	Chloroauric acid 1 mM/plant extract 2.5% (5:1 v/v)80 °C10 minpH NM	Spherical and hexagonal 20 nm	Diffusion37 °C 24 hpH NMInoculum size NMCiprofloxacin 100 µg	E. coli S. aureus	1819 mm	[207]
Inonotus obliquus	Leaves	Chloroauric acid 1 mM/plant extract 5% (19:1 v/v)Room temperature30 min pH NM	Spherical 23 nm	Diffusion37 °C 24 hpH NMInoculum size NMNo control	B. subtilis S. aureus E. coli	121614 mm	[208]
Gloriosa superba	Leaves	Chloroauric acid 1 mM/plant extract 10% (20:1 v/v)50–60 °C10 minpH 5.2	Triangular and spherical20 nm	Diffusion37 °C 24 hpH NM1.5 × 108 CFU/mLAmpicillin 30 µg	B. subtilis ATCC 6633E. coli MTCC 40	6.35.3 mm	[209]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned; NM = not mentioned.

The antimicrobial and antibiofilm properties of AuNPs extend their application to the cosmetic and agricultural fields [184,185]. The applications of antibacterial AuNPs are increasing day-by-day in environmental scenarios, as well as in the impregnation of filters [186,187]. Additionally, AuNPs can bind covalently and non-covalently with secondary coating molecules (e.g., PEG), or other materials, through surface modification. This is to minimize non-specific targeting on other tissues and for the purpose of imaging.

It is of note that previous studies have demonstrated that AuNPs obtained by chemical synthesis are generally not bactericidal, or only weakly so at high concentrations [188,189,190,191]. The reason why these AuNPs may appear to be bactericidal may, inter alia, be due to the bactericidal activity of co-existing organic complexes of Au (I and III) ions that are in the surrounding environment of the AuNPs, and which are not completely removed during centrifugation. The bactericidal properties of AuNPs can be tailored during green synthesis by considering the exposure time required for reduction of Au (III), as well as the speed and number of rounds of centrifugation needed to remove gold ions in excess. This may avoid the observed discrepancies in the antibacterial effects of green AuNPs.

3.2.3. Zinc Oxide Nanoparticles

Zinc is an essential trace element known to have antimicrobial properties against a broad spectrum of microorganisms [210]. The metal zinc is widely present in nature as zinc oxide (ZnO), which is largely used as an antibacterial, antifungal and antibiofilm agent in drug and cosmetic products (e.g., antiseptic powders, shampoos and ointments) [210,211]. To reduce the toxicity of ZnO produced synthetically, a green and eco-friendly synthetic route has been developed to prepare biocompatible ZnO nanoparticles. Table 4 summarizes some green sources alongside the physicochemical characteristics (e.g., size, shape), and the antimicrobial activity of the developed ZnO nanoparticles.

Table 4

Antimicrobial green-synthesized zinc oxide nanoparticles.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics(Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH,Inoculum Density,Positive Control	Tested Bacteria and Fungi	MIC, ZOI or PI *
Cassia alata	Leaves	Zinc acetate 10 mM/plant extract 10% (1:1 v/v)80 °C2 hpH 12	Spherical 60–80 nm	Dilution37 °C24 hpH NM1 × 105 CFU/mLNo control	E. coli	20 μg/mL	[212]
Trifolium pratense	Flowers	Zinc oxide 500 mM/plant extract 2.25% (1:1 v/v)90 °C 24 hpH 6	Hexagonal 60–70 nm	Diffusion 35 ± 1 °C18 h pH NM5 × 105 CFU/mLNo control	S. aureus ATCC 4163E. coli ATCC 25922P. aeruginosa ATCC 6749S. aureus (clinical strain)P. aeruginosa (clinical strain)	3131283129 mm	[213]
Pongamia pinnata	Seed	Zinc acetate 20 mM/plant extract 20% (4:50 v/v)60 °C2 hpH 12	Spherical 30–40 nm	Diffusion37 °C 24 hpH NMInoculum size NMCiprofloxacin 5 μg	P. aeruginosa HQ 693274.1Bacillus licheniformis M235407.1Vibrio parahaemolyticus HQ 693275.1	141712 mm	[214]
Plectranthus amboinicus	Leaves	Zinc nitrate 0.05 mM/plant extract 12% (1:5 v/v)150 °C 6 h pH NM	Hexagonal20–50 nm	Diffusion37 °C24 hpH 7.41 × 105 CFU/mLNo control	S. aureus ATCC 33591	13 mm	[215]
Stevia rebaudiana	Leaves	Zinc acetate 100 mM/plant extract 14% (1:1 v/v)70–80 °C2 hpH NM	Rectangular 10–90 nm	Dilution 37 °C24 h pH NM1.5 × 108 CFU/mLNo control	S. aureus E. coli	22 µg/mL	[216]
Silybum marianum	Seed	Zinc sulfate 1 mM/plant extract 6% (1:50 v/v)37 °C24 hpH 12	Flowers 60 nm	Diffusion37 °C24 hpH NM5 × 106 CFU/mLCefixime **Roxithromycin **	S. aureus ATCC 6538K. pneumoniae ATCC 1705B. subtilis ATCC 6633E. coli ATCC 25922P. aeruginosa ATCC 15442	20 179 10 17 mm	[217]
Linum usitatissimum	Root	Zinc nitrate 0.1 mM/plant extract 10% (1:10 w/v)60 °C3 h pH NM	Hexagonal 35 nm	Diffusion37 °C24 hpH NMInoculum size NMAmoxicillin 10 µg/mL	S. aureus ATCC 6538E. coli ATCC 15224K. pneumoniae ATCC 4619	14 14 12 mm	[218]
Anchusa italic	Flowers	Zinc acetate 100 mM/plant extract 25% (1:10 v/v)70 °C6 hpH NM	Hexagonal 7.6 ± 2.0 nm	Diffusion37 °C24 hpH NM1 × 108 CFU/mLNo control	Bacillus megaterium S. aureus E. coli Salmonella typhimurium	13.614.613 14.4 mm	[219]
Conyza canadensis	Leaves	Zinc nitrate 150 mM/plant extract 6% (1:2 v/v)80 °C20 minpH NM	Spherical 10–50 nm	Diffusion 37 °C24 hpH NMInoculum size NMCiprofloxacin 0.5 mg	E. coli S. aureus	16 14 mm	[220]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned. NM = not mentioned.

3.2.4. Platinum Nanoparticles

Platinum is an inert, biocompatible, nonporous and hypoallergenic metal that is used as an antimicrobial agent for catheters, hip and knee replacement implants, surgical and cardiac stents, implantable cardiovascular defibrillators, etc. [221,222,223]. This metallic chemical element does not corrode into harmful or potentially allergenic substances when kept with soft tissue or bone. Additionally, platinum is not prone to bacterial adhesion and infection since it forms a uniformly smooth surface when plated onto another material, thereby significantly benefiting biomedical applications [221,222,223]. According to the literature, green-synthesized platinum nanoparticles (PtNPs) are found to improve the antibacterial, antifungal and antibiofilm activity of Pt ions (see Table 5).

Table 5

Green platinum nanoparticles exhibiting antimicrobial activity.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics (Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density, Positive Control	Tested Bacteria and Fungi	MIC, DOI or PI *
Garcinia mangostana	Fruit	Hexachloroplatinic acid 1 mM/plant extract 3% (1:1 v/v)50–70 °C15 minpH NM	Spherical20–25 nm	Diffusion35 °C24–48 hpH NM1 × 105 CFU/mLPenicillin G 2 μgMethicillin 5 μgVancomycin 30 μgGentamicin 50 μg Streptomycin 10 μg Ciprofloxacin 5 μgAzithromycin 30 μgClotrimoxazol 25 μg	Staphylococcus spp.Bacillus spp.Pseudomonas spp.Klebsiella spp.	100 1211 mm	[224]
Citrus sinensis	Peel	Hexachloroplatinic acid 10 mM/plant extract 10% (9:1 v/v)80 °C24 hpH NM	Spherical50 nm	Diffusion30 ± 1 °C24 hpH 4Inoculum size NMNo control	Aeromonas hydrophila	4 mm	[225]
Sechium edule	Fruit	Platinum (II) chloride 1 mM/plant extract 12.5% (1:1 v/v)100 ± 5 °C12 hpH 9	Spherical 28 nm	Diffusion37 °C24 hpH NM5 × 105 CFU/mL Ciprofloxacin 30 μg Cefprozil 30 μg	B. subtilis E. coli	2524 mm	[226]
Spinacia oleracea	Leaves	Hexachloroplatinic acid 20 mM/plant extract 75% (2:1 v/v)100 °C24 hpH NM	Rod 154 nm	Diffusion37 °C24 hpH NM1 × 105 CFU/mLNo control	S. typhi MTCC 098	13 mm	[227]
Taraxacum laevigatum	Powder	Hexachloroplatinic acid 10 mM/plant extract 5% (5:1 v/v)90 °C10 minpH NM	Spherical 2–7 nm	Diffusion 37 °C24 hpH NM5 × 105 CFU/mLStreptomycin **	B. subtilis P. aeruginosa	1815 mm	[228]
Cerbera manghas	Leaves	Hexachloroplatinic acid 1 mM/plant extract 2% (19:1 v/v)25 °C2 hpH NM	Spherical 9–12 nm	Diffusion 37 °C24 hpH NM1 × 108 CFU/mLStreptomycin 0.25 mg/mL	Vibrio cholerae S. aureus Streptococcus pyogenes S. typhi E. coli	2019131211 mm	[229]
Prunus yedoensis	Gum	Hexachloroplatinic acid 100 mM/plant extract 25% (5:1 v/v)80 °C5 hpH NM	Spherical and oval10–50 nm	Diffusion37 °C 48 h pH NMInoculum size NMNystatin **	Phytophthora capsici Phytophthora drechsleri Didymella bryoniae Colletotrichum acutatum Cladosporium fulvum	0001518 mm	[230]
Curcuma longa	Seed	Hexachloroplatinic acid 1 mM/plant extract 1% (1:1 v/v)80 °C2 hpH 10	Spherical 9 nm	Dilution 37 °C3 hpH NM1 × 108 CFU/mLNo control	E. coli CD-496E. coli CD-2E. coli CD-3E. coli CD-19E. coli CD-549S. aureus CD-1578MRSA CD 489	16161616166432 nM	[231]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned. NM = not mentioned; MRSA = methicillin-resistant S. aureus.

3.2.5. Palladium Nanoparticles

Palladium (Pd) is a noble metal widely used as a platinum substitute due to its similar attributes and functionalities [203]. Indeed, Pd’s inherent biocompatibility, hypoallergenicity, chemical inertness, non-porosity and antimicrobial potential make it valuable for the manufacture of medical devices, thereby preventing corrosion and disease infections. Biogenic PdNPs also show outstanding antimicrobial properties (see Table 6); for this reason, they are widely applied in dental and surgical implants as well as prostheses [221].

Table 6

Green palladium nanoparticles with antimicrobial activity.

Plant Type	Part Used	Operative Conditions for Synthesis	NpCharacteristics(Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density, Positive Control	Tested Bacteria and Fungi	MIC, DOI or PI *
Moringa oleifera	Peel	Palladium acetate 10 mM/plant extract 10% (4:1 v/v)80 °C5 minpH NM	Spherical 27 nm	Diffusion 37 °C24 hpH NMInoculum size NMAmoxicillin **	S. aureus E. coli	11 mm	[232]
Prunus yedoensis	Leaves	Palladium chloride 1 mM/plant extract 25% (9:1 v/v)80 °C30 minpH NM	Spherical 50–150 nm	Diffusion 37 °C24 hpH NMInoculum size NMAmoxicillin **	B. subtilis P. aeruginosa	65 mm	[230]
Cissus quadrangularis	Stem	Palladium chloride 0.05 mM/plant extract 10% (1:5 v/v)37 °C10 minpH NM	Spherical 12–26 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	E. coli	17 mm	[233]
Camellia sinensis	Leaves	Palladium chloride 1 mM/plant extract 1% (1:1 v/v)40 °C30 minpH NM	Spherical6–18 nm	Diffusion 37 °C24 hpH NM1 × 108 CFU/mLStreptomycin **	S. epidermidis S273E. coli E266	1714 mm	[234]
Garcinia pedunculata	Leaves	Palladium acetate 1 mM/plant extract 20% (2:1 v/v)121 °C15 minpH NM	Spherical 2–4 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	Cronobacter sakazakii AMD04	0.3 mm	[235]
Phoenix dactylifera	Leaves	Palladium chloride 3 mM/plant extract 10% (5:1 v/v)37 °C10 minpH NM	Spherical2–5 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	P. aeruginosa	26 mm	[236]
Arabidopsis thaliana	Leaves	Palladium chloride 5 mM/plant extract 1% (10:1 v/v)80 °C24 hpH NM	Spherical 20–40 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	S. aureus	29 mm	[237]
Acacia senegalensis	Gum	Tetrachloropalladic acid 1 mM/plant extract 0.2% (1:1 v/v)100 °C6 hpH NM	Spherical 10 nm	Diffusion37 °C24 hpH NM0.5 × 105 CFU/mLNo control	Bacillus cereus S. aureus Streptococcus agalatiae	181617 mm	[238]
Bauhinia variegate	Bark	Palladium chloride 1 mM/plant extract 10% (4:1 v/v)60 °C30 minpH NM	Irregular 2–9 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	B. subtilis MTCC 441S. aureus MTCC 737E. coli MTCC 1687C. albicans MTCC 183	16617 mm	[239]
Allium cepa	Bulb	Palladium chloride 10 mM/plant extract 10% (1:5 v/v)100 °C2 hpH NM	Spherical 19 nm	Diffusion 37 °C24 hpH NM1 × 106 CFU/mLNo control	Bacillus cereusS. aureusMicrococcus spp.E. coli Klebsiella spp.Proteus spp.	362740221817 mm	[240]
Filicium decipiens	Leaves	Palladium chloride 1 mM/plant extract 10% (9:1 v/v)37 °C96 hpH NM	Spherical 2–22 nm	Diffusion 37 °C24 hpH NM1 × 105 CFU/mLLevofloxacin **	B. subtilis S. aureus E. coli P. aeruginosa	12122724 mm	[241]
Phyllanthus emblica	Seed	Palladium acetate 870 mM/plant extract 10% (4:1 v/v)60 °C3 hpH NM	Spherical 28 nm	Diffusion 37 °C24 hpH NMInoculum size NMStreptomycin 50 µg/mL	B. subtilis S. aureus P. aeruginosa Proteus mirabilis	8.98.27.64.3 mm	[242]
Eucommia ulmoides	Bark	Palladium chloride 10 mM/plant extract 20% (5:1 v/v)80 °C30 minpH 6	Spherical 2 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	S. aureus E. coli	2529 mm	[243]
Delonix regia	Leaves	Palladium chloride 0.5 mM/plant extract 25% (9:1 v/v)28 °C3 hpH NM	Spherical 2–4 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	Streptococcus mitis	12 mm	[244]
Coriandrum sativum	Seed	Palladium chloride 10 mM/plant extract 5% (1:5 v/v)60 °C3 hpH NM	Spherical113 nm	Diffusion 37 °C18 hpH NM5 × 108 CFU/mLStreptomycin 50 µg/mL	S. aureus E. coli Salmonella enterica	138.510 mm	[245]
Piper betle	Leaves	Palladium chloride 1 mM/plant extract 20% (1:10 v/v)30 °C1 hpH NM	Spherical4 nm	Diffusion30 °C 48 h pH NMInoculum size NMClotrimazol 1 mg/mL	A. niger	34 mm	[246]
Rosmarinus officinalis	Leaves	Palladium acetate 100 mM/plant extract 10% (2:1 v/v)37 °C24 hpH NM	Spherical15 nm	Diffusion 37 °C20 hpH NM1 × 106 CFU/mL Ciprofloxacin **	S. aureus E. coli S. epidermidis Micrococcus luteus	24252120 mm	[247]
				Difusion 32 °C1 week pH NM1 × 106 CFU/mL Nystatin **	C. albicans Candida parapsilolis Candida glabrata Candida krusei	0000 mm
Boswellia serrata	Gum	Palladium chloride 1 mM/plant extract 0.5% (1:1 v/v)121 °C30 minpH NM	Spherical 2–9 nm	Diffusion37 °C24 hpH NM1 × 106 CFU/mLGentamicin 10 µg	S. aureus ATCC 25923P. aeruginosa ATCC 27853	2628 mm	[248]
Coffea arabica	Powder	Palladium chloride 100 mM/plant extract 8% (1:5 v/v)60 °C3 hpH NM	Spherical 20–60 nm	Diffusion 37 °C24 hpH NM2 × 108 CFU/mLAmpicillin **	Enterococcus faecalis S. typhi S. epidermidis	121212 mm	[249]
Morus alba	Fruit	Palladium chloride 2 mM/plant extract 20% (5:1 v/v)80 °C3 hpH NM	Spherical and non-regular50–150 nm	Diffusion 37± °C 24 hpH NM1.5 × 108 CFU/mLAmoxicillin **	Listeria monocytogenes ATCC 19115E. coli O157:H7	2629 mm	[120]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned. NM = not mentioned.

3.2.6. Other Green Metallic Nanoparticles

Green CuO, Fe2O3, Fe3O4, NiO, MgO, MnO, Mn5O8 and TiO2 nanoparticles are also found to be effective against bacteria and fungi (see Table 7 and Table 8) [25,250]. The inorganic nanoparticles conjugated with nanocellulose-based antimicrobial materials may have huge potential applications in the area of drug delivery, (bio)pharmaceuticals, (dermo)cosmetology, wound dressing, tissue engineering, food packaging, water treatment, air filtration, coating, mask cartridges, etc. [107,115,119]. Nevertheless, microbiological studies remain scarce for this type of nanoparticle, and further investigations are needed.

Table 7

Green copper nanoparticles exhibiting antimicrobial activity.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics(Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density, Positive Control	Tested Bacteria and Fungi	MIC, DOI or PI *
Cymbopogon citratus	Leaves	Copper (II) sulfate 0.25 mM/plant extract 50% (2:1 v/v) 60 °C3 h pH 12	Spherical, hexagonal and oval12–14 nm	Diffusion 37 °C24 hpH NM1 × 107 CFU/mLNo control	E. coli ESβL-336MSSAMRSA	201816 mm	[251]
Ziziphus spina-christi	Fruits	Copper (II) sulfate 20 mM/plant extract 6% (1:10 v/v)80 °C1 hpH NM	Spherical 9 nm	Diffusion 37 °C24 hpH NMInoculum size NMNo control	E. coli S. aureus	22 mm	[252]
Enicostemma axillare	Leaves	Copper (II) sulfate 5 mM/plant extract 10% (10:1 v/v)50 °C24 h pH 7	Spherical 44 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	E. coli P. aeruginosa K. pneumoniae S. aureus Proteus vulgaris	468118 mm	[253]
Phyllanthus emblica	Fruits	Copper (II) sulfate 20 mM/plant extract 50% (3:1 v/v)80 °C15 min pH 10	Flakes 15–30 nm	Diffusion37 °C24 hpH NMInoculum size NMCiprofloxacin 25 μg	E. coli S. aureus	1414 mm	[254]
Carica papaya	Leaves	Copper (II) sulfate 5 mM/plant extract 10% (9:1 v/v)60 °C10 minpH NM	Rod 40 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	E. coli S. aureus P. aeruginosa	9910 mm	[255]
Sida acuta	Leaves	Copper (II) sulphate 1000 mM/plant extract 4% (2:1 v/v)100 °C5–7 hpH NM	Spherical 50 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	E. coli Proteus vulgaris	1511 mm	[256]
Prosopis cineraria	Leaves	Copper (I) acetate 5 mM/plant extract 10% (1:1 v/v)Room temperature24 hpH NM	Hexagonal19–32 nm	Diffusion37 °C24 hpH NMInoculum size NMCefotaxim **	Proteus vulgaris P. aeruginosa K. pneumoniae E. coli S. aureus S. epidermidis	171822221923 mm	[257]
Syzygium aromaticum	Buds	Copper (II) acetate 1 mM/plant extract 100% (5:1 v/v)30 °C15 min pH NM	Spherical 20 nm	Diffusion37 °C24 h pH NMInoculum size NMNo control	Staphylococcus spp.E. coli Pseudomonas spp.Bacillus spp.	5678 mm	[258]
				Diffusion37 °C72 h pH NMInoculum size NMNo control	A. nigerA. flavusPenicillium spp.	556 mm
Ruellia tuberosa	Leaves	Copper (II) sulfate 1 mM/plant extract 5% (1:1 v/v)100 °C7–8 h pH NM	Nanorod20–100 nm	Diffusion37 °C24 hpH NMInoculum size NMStreptomycin **	S. aureus K. pneumoniae E. coli	131418 mm	[259]
Punica granatum	Peels	Copper (II) sulfate 50 mM/plant extract 10% (1:1 v/v)80 °C10 minpH NM	Spherical 15–20 nm	Diffusion37 °C 24 hpH NMInoculum size NMStreptomycin **	P. aeruginosa MTCC 424 Salmonella enterica MTCC 1253 Micrococcus luteus MTCC 1809Enterobactera erogenes MTCC 2823	192020 19 mm	[260]
Asparagus adscendens	Leaves	Copper (II) sulfate 1 mM/plant extract 5% (10:1 v/v)100 °C1 hpH NM	Spherical 10–15 nm	Diffusion 37 °C24 hpH NM1 × 108 CFU/mLAmpicillin 25 µg/mL	E. coli B. subtilis S. typhi K. pneumoniae S. aureus	2018211817 mm	[261]
Gloriosa superba	Leaves	Copper (II) sulfate 1 mM/plant extract 5% (4:1 v/v)60 °C3–4 minpH NM	Spherical 5–10 nm	Diffusion37 °C24–36 hpH NMInoculum size NMCiprofloxacin 0.5 µg/µL	Klebsiella aerogenes NCIM 2098E. coli NCIM 5051S. aureus NCIM 5022Pseudomonas desmolyticum NCIM 2028	151365 mm	[262]
Cassia auriculata	Leaves	Copper (II) sulfate1 mM/plant extract 5% (4:1 v/v)Room temperature5 hpH NM	Clusters38 nm	Diffusion37 °C24 hpH NM1 × 108 CFU/mLAmoxicillin **	E. coli P. aeruginosa S. aureus Proteus mirabilis Bacillus cereus K. pneumoniae	161014161814 mm	[263]
Bersama abyssinica	Leaves	Copper (I) acetate 100 mM/plant extract 10% (1:1 v/v)80 °C2 hpH NM	Spherical31 nm	Diffusion37 °C24 hpH NMInoculum size NMGentamicin **	S. aureus B. subtilis E. coli P. aeruginosa	126146 mm	[264]
Datura innoxia	Leaves	Copper (II) sulfate 1 mM/plant extract 5% (1:1 v/v)100 °C1 hpH NM	Nanoclusters90–200 nm	Diffusion37 °C24 hpH NMInoculum size NMPlantomycin *	Xanthomos oryzae pv. oryzae	24 mm	[265]
Zingiber officinale	Rhizome	Copper (II) sulfate 5 mM/plant extract 30% (5:3 v/v)60 °C4 hpH NM	Spherical31 nm	Diffusion37 °C24 hpH NMInoculum size NMCiprofloxacin **	E. coli	22 mm	[266]
Vaccinium arctostaphylos	Fruit	Copper (II) acetate/plant extract 5% (1:20 w/v) 60 °C24 h pH 10	Spherical 14 nm	Diffusion37 °C24 hpH NMInoculum size NMNitrofurantoïn **	E. coli	22 mm	[267]
Cissus arnotiana	Leaves	Copper (II) sulphate 10 mM/plant extract 1% (9:1 v/v)60 °C4 hpH NM	Spherical 60–90 nm	Diffusion37 °C18 hpH NMInoculum size NMAmpicillin **	E. coliStreptococcus spp.Rhizobium spp.Klebsiella spp.	2220.216.318.3 mm	[268]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned. NM = not mentioned; MRSA = methicillin-resistant S. aureus; MSSA = methicillin-susceptible S. aureus.

Table 8

Antimicrobial green-synthesized iron nanoparticles.

Plant Type	Part Used	Operative Conditions for Synthesis	NP Characteristics (Shape and Size)	Microbiological Analyzes (Operative Conditions)			Refs.
				Methods, Incubation Temperature, Incubation Time, pH, Inoculum Density,Positive Control	Tested Bacteria and Fungi	MIC, DOI or PI *
Withania coagulans	Berries	Iron (III) chloride 2000 mM/plant extract 12% (5:1 v/v)90 °C30 minpH NM	Rod 16 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	P. aeruginosa S. aureus	2423 mm	[269]
Acacia nilotica	Pods	Iron (II) sulfate 100 mM/plant extract 10% (3:2 v/v)Room temperature24 hpH 6	Irregular39 nm	Diffusion30 °C24 hpH NMInoculum size NMNo control	E. coliMRSAS. typhiS. aureusC. albicans	1724232525 mm	[270]
Musa ornate	Flowers	Iron (III) chloride 1 mM/plant extract 10% (1:1 v/v)70–80 °C1 hpH NM	Spherical 20–40 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	Streptococcus agalactiae S. aureus Salmonella enterica E. coli	2832 00 mm	[271]
Skimmia laureola	Leaves	Iron (III) chloride 100 mM/plant extract 10% (1:1 v/v)Room temperature30 minpH NM	Spherical 56–350 nm	Diffusion37 °C24 hpH NMInoculum size NMStreptomycin 200 ppm	Ralstornia solanacearum	18 mm	[272]
Lagenaria siceraria	Leaves	Iron (III) chloride 10 mM/plant extract 5% (1:1 v/v)40 °C60 minpH NM	Cubic 30–100 nm	Diffusion37 °C24 hpH NMInoculum size NMAmpicillin 20 mg/mL	S. aureus E. coli	1417 mm	[273]
Trigonella foenum-graecum	Seed	Iron (II) chloride 1000 mM/plant extract 5% (1:2 v/v)Room temperature2 hpH 10	Spherical ∼20 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	E. coli ATCC 11775S. aureus ATCC 6538	2224 mm	[148]
Dodonaea vicosa	Leaves	Iron (III) chloride 10 mM/plant extract 20% (2:1 v/v) 50 °C1 hpH NM	Spherical50–60 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	E. coli MTCC 443K. pneumoniae NCIM 2079B. subtilis MTCC 441S. aureus MTCC 4032Pseudomonas fluorescens MTCC 121	8101214 24 mm	[274]
Couroupita guianensis	Peel	Iron (III) chloride 100 mM/plant extract 5% (1:1 v/v)80 °C30 minpH 10	Spherical 7–80 nm	Diffusion37 °C24 hpH NM1 × 105 CFU/mLStreptomycin 1 mg	S. aureus MTCC 96E. coli MTCC 2939S. typhi MTCC 3917K. pneumoniae MTCC 530	8151512 mm	[275]
Psidium guajava	Fruit	Iron (III) chloride 500 mM/plant extract 4% (4:1 v/v)100 °C1 h pH NM	Hexagonal 27 nm	Diffusion37 °C18–24 hpH NMInoculum size NMGentamicin 10 μg	Bacillus cereus E. coli K. pneumoniae S. aureus	14171014 mm	[276]
Punica granatum	Peel	Iron (III) chloride 150 mM/plant extract 4.6% (5:2 v/v) 20 °C5 hpH NM	Spherical 20–90 nm	Diffusion30 °C24 hpH NMInoculum size NMStreptomycin **	P. aeruginosa	22 mm	[277]
Argemone mexicana	Leaves	Iron (III) chloride 25 mM/plant extract 3% (1:1 v/v) 45 °C12 hpH NM	Spherical10–30 nm	Diffusion37 °C24 hpH NM1 × 106 CFU/mLStreptomycin 30 µg	E. coli MTCC 443B. subtilis MTCC 425Proteus mirabilis MTCC 441	131810 mm	[278]
Ruellia tuberosa	Leaves	Iron (II) sulphate 1000 mM/plant extract 5% (1:1 v/v)80 °C30 minpH NM	Hexagonal 53 nm	Diffusion37 °C24 hpH NM1 × 106 CFU/mLStreptomycin **	K.pneumoniae E. coli S.aureus	131611 mm	[279]
Leucas aspera	Leaves	Iron (III) chloride 5 mM/plant extract 20% (1:1 v/v)80 °C15 minpH NM	Irregular rhombic117 nm	Diffusion37 °C24 hpH NMInoculum size NMAmpicillin 10 µg	E. coli K. pneumoniae Proteus mirabilis Salmonella enterica Shigella flexneri Vibrio cholera P. aeruginosa Bacillus cereus S. aureus Listeria monocytogens	10101119221020001112 mm	[280]
Eichhornia crassipes	Leaves	Ferrous sulphate 100 mM/plant extract 5% (1:1 v/v)55 °C2 hpH 10	Rod10–100 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	S. aureus Pseudomonas fluorescens E. coli	232320 mm	[281]
Sida cordifolia	Whole the plant	Iron nitrate 10 mM/plant extract 5% (2:1 v/v)60 °C5 minpH NM	Spherical 10–22 nm	Diffusion37 °C24 hpH NMInoculum size NMNo control	B. subtilis S. aureus E. coli K. pneumoniae	17151519 mm	[282]
Trigonella foenum-graecum	Seed	Iron (III) chloride 10 mM/plant extract 0.04% (1:20 v/v)30 °C15 minpH NM	Spherical 11 nm	Diffusion37 °C24 hpH NMInoculum size NMStreptomycin **	E. coli S. aureus	2219 mm	[283]
Piliostigma thonningii	Flowers	Iron (II) chloride 1 mM/plant extract 20% (9:1 v/v)80 °C2 minpH NM	Spherical 20–100 nm	Diffusion37 °C24 hpH NMInoculum size NMGentamycin **	E. coli S. aureus	2120 mm	[284]

* MIC = minimal inhibition concentration; ZOI = zone of inhibition; PI = percentage of inhibition. ** The quantity or concentration is not mentioned. NM = not mentioned; MRSA = methicillin-resistant S. aureus.

Considering the above, it is clear that green MNPs are promising as antimicrobials. They can solve many problems in medicine, human health, nanomedicine and other fields (Figure 6).

Figure 6

Overview of the different applications of antimicrobial MNPs.

4. Methods for Testing Antibacterial and Antifungal Activities of Green MNPs

Planktonic bacterial and fungal cells are free-living or free-floating bacteria and fungi that are responsible for several infectious diseases. The biological properties of green-synthesized MNPs against bacteria and fungi are well-documented in the literature [285,286,287,288,289,290,291,292]. To assess these antimicrobial potentials, different physical and analytical characterization techniques can be used [293,294]. Physical characterization techniques (e.g., atomic force microscopy, fluorescence spectroscopy, ultra-microtome-based transmission electron microscopy, inductively coupled plasma mass spectroscopy) have been developed to ascertain different information related to the interactions between MNPs and microorganisms [295]. They also provide information about the microbial killing mechanisms of green MNPs [296,297]. However, these physical techniques are beyond the scope of the present review.

Analytical techniques (e.g., colony-forming-unit (CFU) assay, live-dead staining assay, disk diffusion assay, minimum inhibitory concentration assay, capillary electrophoresis, enzyme-linked immunoassay, and polymerase chain reaction) constitute the most common methods used for testing the antimicrobial activity of MNPs [298]. As shown in Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7, diffusion and dilution susceptibility testing represent the main analytical techniques for the assessment of the antimicrobial activity of MNPs [199,247,299,300,301,302]. Both these analytical techniques are described below.

4.1. Analytical Techniques: Diffusion and Dilution Susceptibility Testing Methods

In diffusion susceptibility testing methods (also known as disk diffusion assay), green-synthesized MNPs impregnated in discs diffuse in an agar medium (which contains the tested bacterium) and surround the discs. This diffusion (or spread) phenomenon leads to inhibition of the growth of bacteria and fungi in an area around the discs [303]. The diameter of this inhibition zone is determined by the distance that the inhibitory concentration of the green-synthesized MNPs may travel before a certain microbial density [304]. Antibiograms and antifungiograms obtained by this method are widely used because of their simplicity, low-cost, ability to test a large number of microorganisms and antimicrobial agents, and readily interpretable results [305]. However, this method is not suitable for determining the minimum inhibitory concentration (MIC), i.e., the minimum concentration of biofabricated MNPs that inhibit the visible growth of microorganisms after a given time (18–24 h) of incubation at a predefined temperature [306]. To determine the MIC by diffusion methods, some researchers impregnate sterile paper discs with a volume of MNP solutions, while others cut wells of a certain diameter (generally 6 mm) in fungi- or bacteria-containing plates using a sterile corkborer, and then add a volume of nanoparticle solutions into each well [307,308,309].

In dilution susceptibility testing methods (so-called dilution methods), the evaluation of antimicrobial activity of eco-friendly MNPs can be performed using either agar culture medium (e.g., Mueller–Hinton agar (MHA)) (agar dilution method) or liquid culture medium (e.g., Mueller–Hinton broth (MHB)) (broth or liquid dilution method) [310,311,312,313]. Dilution methods are more suitable for determining the MIC than diffusion methods, as they provide a better quantitative estimate of the antibacterial and antifungal activity of MNPs [314,315]. The agar dilution method requires mixing of different concentrations of nanoparticle solutions in the molten MHA medium. After pouring and solidifying the obtained mixture into Petri dishes on a level surface to a specific agar depth, a quantity of standardized bacterial or fungal suspension is inoculated on the agar surface by multiple streaks [316]. The agar plates are then incubated according to validated procedures and/or practice guidelines of accredited organisms, such as the European Committee on Antimicrobial Susceptibility Testing (EUCAST) and the Clinical and Laboratory Standards Institute (CLSI) [317,318].

On the other hand, the MHB dilution method can be performed in tubes (macromethod or macrodilution method) or in microtiter plates (micromethod or microdilution method) [319]. The broth microdilution method is preferred over the broth macrodilution method for the development of antibiograms because it is easy to handle, cost-effective and can be automated for the preparation of nanoparticle dilutions and the reading of the MIC [303,316,320,321].

Although all the results presented in Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7 confirm the reliability of the diffusion and dilution methods for the MNP susceptibility testing of different microbes, only a very limited number of studies have compared the MIC values resulting from the dilution method with the MIC values (or DOIs) obtained by the diffusion method, or other methods, such as the E-test. The objective would be to identify the most feasible, quantitative and cost-effective methods for high-throughput MNP susceptibility testing that can be used in any laboratory setting and for any clinical bacteria and fungi. With respect to future studies, the comparison of data from dilution versus diffusion methods will likely guide the reader in choosing which is most appropriate for particular microorganisms, experimental conditions or biomedical applications.

4.2. Factors Influencing the Evaluation of Antimicrobial Activities of Metallic Nanoparticles

Factors that can influence either the diameter of the inhibition zone or the MIC, and modify the antibacterial and antifungal activities of MNPs include: (i) the types and species of bacteria and fungi, (ii) the diversity of strains within bacterial and fungal species, (iii) the inoculum density, (iv) the composition and pH of the culture medium, (v) the disc application timing, (vi) the temperature and time of incubation, (vii) the depth of the agar medium, (viii) the spacing of the antibiotic discs, (ix) the nature and concentration of the capping and stabilizing agents, (x) the size, shape and zeta potential of nanoparticles, (xi) the type, content and quality control of antibiotic discs, (xii) the impregnation of nanoparticles in sterile blank discs, and (xiii) the additive and synergistic effects of antimicrobial nanoparticles in combination with conventional or non-traditional antibiotics [303,321,322,323]. The impact of some of these parameters on the antimicrobial assessment of MNPs is discussed in the following paragraphs.

4.2.1. Types of Bacterial and Fungal Species and Strains

The human body hosts a whole community of commensal, saprophytic and pathogenic microorganisms grouped under the term “microbiota” [324,325].

The skin or flora microbiota can be resident or transient. The resident bacteria are dominated by commensal bacteria (e.g., Staphylococcus epidermidis, Staphylococcus warneri) and coryneform bacteria (e.g., Brevibacterium epidermidis, Arthrobacter globiformis, Corynebacterium spp.). In contrast, the transient microbiota are composed of saprophytic microorganisms (e.g., Propionibacterium acnes) and opportunistic pathogenic bacteria (e.g., Staphylococcus aureus, Pseudomonas aeruginosa, Bacillus spp., Treponema pallidum, Acinetobacter johnsonii and Acinetobacter baumannii) [326,327]. Primary cutaneous mold infections are predominantly caused by Aspergillus flavus, Aspergillus niger, Fusarium spp., Mucor spp., Rhizopus stolonifer, Malassezia globosa, Trichosporon cutaneum, Alternaria alternata, Candida albicans, Torulopsis spp., Trichosporon cutaneum, Penicillium spp., etc. [326,327,328,329].

The oral flora is largely dominated by anaerobic bacterial genera, such as Actinomyces, Bacteroides, Lactobacillus, Leptotrichia, Treponema, Streptococcus and Peptococcus. Several kinds of microfungi, such as Aspergillus, Candida, Cladosporium, Cryptococcus, Fusarium and Penicillium are also found in the mouth [330].

In the small intestine and the colon, the intestinal microbiota is mostly represented by Escherichia coli, Helicobacter pylori, Pseudomonas aeruginosa (bacteria), Candida albicans and Saccharomyces boulardii (fungi) [331].

Despite the high number of (opportunistic) pathogenic microorganisms, most research papers are limited to assessing the antimicrobial susceptibility of three bacterial species (Staphylococcus aureus, Escherichia coli and Pseudomonas aeruginosa) and three fungal isolates (Aspergillus flavus, Aspergillus niger and Candida albicans) (Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7; Figure 7) [9,192,332,333,334,335,336,337]. However, testing the antibacterial and antifungal susceptibility of a large number of pathogenic microorganisms from cutaneous, oral and intestinal microbiota would help to expand the spectrum of activity of biosynthesized MNPs. Additionally, although most studies attest that Gram-negative bacteria are more sensitive to antimicrobial nanoparticles than Gram-positive bacteria [338,339,340,341,342], it is important to always confirm this rule with a broader range of green MNPs, as well as bacterial and fungal species and strains. By doing so, the chances of detecting certain exceptions increases, as previously reported [9].

Figure 7

(A) Percentage of bacteria citations (n = 2065). Others (8.8%) include ca. 70 bacterial species, such as Acinetobacter baumannii, Bacillus megaterium, Enterobacter cloacae, Klebsiella planticola, Pseudomonas putida, Staphylococcus saprophyticus and Shigella dysenteriae. (B) Percentage of fungi citations (n = 326). Others (20.0%) include ca. 15 fungal species, such as Aspergillus terreus, Candida krusei, Candida freundii, Fusarium oxysporum, Penicillium italicum, Penicillium notatum and Phanerochaete sordida.

Most of the microbiological strains that have been tested for their susceptibility against green-synthesized metallic nanoparticles were procured from the American Type Culture Collection (ATCC) [121,343,344,345], the Microbial Type Culture Collection (MTCC) [274,346,347,348], or the National Collection of Industrial Microorganisms (NCIB) [274,334,348] (Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7, Figure 8). However, laboratory strains of the same bacterial and fungal species may show differences in sensitivity against MNPs [349]. This is particularly the case for β-lactamase-negative and β-lactamase-positive laboratory strains (e.g., E. coli ATCC 25922 vs. 35218, and S. aureus ATCC 25923 vs. 38591). Therefore, it is recommended that the ATCC, MTCC or NCIB strain numbers of all microorganisms are recorded. Moreover, it would be desirable to assess and compare the antimicrobial activities of green MNPs on these different strains of bacterial and fungal species.

Figure 8

(A) Percentage of microbiological analysis citations (n = 635). (B) Percentage of inoculum size citations (n = 635). (C) Percentage of microbial strain source citations (n = 2391). Others (5.4%) include CMCC (China Medical Culture Collection), KACC (Korean Agricultural Culture Collection), KCCM (Korean Culture Center of Microorganisms) and PTCC (Persian Type Culture Collection) (NM* = not mentioned). (D) Percentage of positive control citations (n = 752).

It is worth noting that microbial strains from laboratory and environmental sources generally have lower virulence than their corresponding clinical strains [350,351,352,353]. Consequently, discrepancies may be observed in the antimicrobial activity of MNPs against both clinical and laboratory strains. Hence, to better fight nosocomial infections and prevent the emergence of newly emerging clinical strains and multi-drug-resistant bacteria (e.g., methicillin-resistant S. aureus, multi-drug-resistant P. aeruginosa, etc.), pharmaceutical strategies based on the development of MNPs, and the evaluation of their antimicrobial activity, should include an even greater number of bacterial and fungal pathogens.

4.2.2. Inoculum Density

CLSI and EUCAST consider inoculum density to be an important variable in susceptibility testing [354,355]. Indeed, if the inoculum is too large, the chances of reducing the diameter of inhibition zones (or enhancing the MIC) of the studied microorganisms by MNPs increases [354,355,356,357,358,359]. The consequence is that sensitive strains can be considered to be relatively resistant when they are not. Conversely, if the inoculum is too small, the zone of inhibition can be enhanced and the MIC reduced, thereby relatively resistant strains can be considered to be sensitive or susceptible [360].

According to American, European and Japanese pharmacopeias [361,362,363], pleasing results can be obtained with an inoculum size of ca. 108 CFU/mL. In contrast, the CLSI states that inoculum size ranging from 2 × 105 to 8 × 105 CFU per mL (or 2 × 103 to 10 × 103 viable cells per spot of 10 µL) can produce confluent growth, thereby leading to optimal results (Table 9) [364,365]. However, referring to most published data related to the microbiological analysis of green MNPs, the initial inoculum sizes of tested microorganisms varies from 104 to 1.5 × 108 CFU/mL [195,235,301,366,367]. Regrettably, we note that some research groups do not mention this density at all (Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7 and Figure 8).

Table 9

Experimental conditions for antimicrobial susceptibility testing methods as recommended by CLSI (Adapted from [303,318,365,368,369,370,371,372]).

Methods	Microorganism	Growth Medium	Final Inoculum Size	IncubationTemperature	IncubationTime 1
Disk diffusion	Bacteria	MHA	1–2 × 108 CFU/mL	35 ± 2 °C	16–18 h
	Fungi	MHA + GMB (Yeast)	1–5 × 106 CFU/mL (yeast)	35 ± 2 °C	20–24 h
		Non-supplemented MHA (molds)	0.4–5 × 106 CFU/mL (molds)	–	–
Broth dilution	Bacteria	MHB	5 × 105 CFU/mL	35 ± 2 °C	20 h 2
	Fungi	RPMI 1640 (yeast)	0.5–2.5 × 103 CFU/mL (yeast)	35 °C	24–48 h (yeast) 3
		RPMI 1640 (molds)	0.4–5 × 104 CFU/mL (molds)	35 °C	48 h (molds) 4
Agar dilution	Bacteria	MHA	1 × 104 CFU/spot	35 ± 2 °C	16–20 h
Time-kill test	Bacteria	MHB	5 × 105 CFU/mL	35 ± 2 °C	0, 4, 18 and 24 h

1 MHA: Mueller–Hinton agar; MHB: Mueller–Hinton broth; GMB: glucose (2%) and methylene blue (0.5 mg/mL); RPMI 1640: Roswell Park Memorial Institute medium. 2 The USP recommends incubating Escherichia coli, Pseudomonas aeruginosa and Staphylococus aureus at 32.5 ± 2.5 °C for 18–24 h. In contrast, Candida albicans should be incubated at 22.5 ± 2.5 °C for 44–52 h, and Aspergillus niger at 22.5 ± 2.5 °C for 6–10 days. 3 24–48 h for microdilution and 46–50 h for macrodilution.4 48 h for both microdilution and macrodilution.

Hence, to avoid false-positive or false-negative test results (also known as the inoculum effect) [364], the inoculum density must be standardized. To carefully determine the size of the inoculum and to perform colony counts, McFarland standards or calibrators should be used; 0.5 McFarland standards correspond to approximately 1–2 × 108 CFU/mL [358]. The obtained inoculums need to be used within 15 min of preparation in order to avoid the premature growth of microorganisms [358,365].

4.2.3. Agar Depth and Spacing of Impregnated Discs

Among the parameters that can affect the antimicrobial activity of MNPs, the depth of the agar medium and the spacing of discs impregnated with nanoparticles can also be cited. Indeed, a reduced inhibition zone may be obtained on very thick media, while the reverse is true for media that are too thin [373]. To counteract these phenomena, the depth of the agar medium should be between 4 and 10 mm [374,375]. However, it is difficult to know if this recommendation is always applied or not, especially since the agar depth is not mentioned in most of the studies pertaining to MNPs with antimicrobial properties. To ensure a better comparison of data from one study to another, subsequent research should respect recommendations based on the depth of the agar medium.

It is suggested to place a maximum of seven discs (of 10 mm) on 90–100 mm diameter plates to avoid the overlap of different inhibition zones and/or their deformation near the edge of the plates [358,376,377,378]. This reduces artifacts and the difficulty of comparing microbiological data from one study to another. Once again, very little information on agar depth and the spacing of discs can be obtained from the literature related to MNPs with antimicrobial properties. Nevertheless, based on some data available in the literature, it appears that this provision is not respected in many studies.

4.2.4. Timing of Disc Application

The timing for application of discs impregnated with MNPs on the agar medium (diffusion method) and the dilution of nanoparticle solutions in the microorganism suspensions in glass test tube dilution (macrodilution method), or microtiter plastic plates containing 96 wells (microdilution method), are also considered as major variables that can influence the antibacterial and antifungal activities of MNPs [358].

Based on CLSI and EUCAST guidelines, disc application and dilution operations should be performed within 15 min following inoculum preparation [359]. Indeed, if the plates, tubes and microplates seeded with the tested microorganisms are left at room temperature for periods longer than the standard time, microorganisms can start growing before the application of discs or dilutions of nanoparticle solutions. By reducing the diameter of the inhibition zone and/or increasing the MIC values, these phenomena may result in a susceptible strain being wrongly reported as resistant [360].

4.2.5. Temperature and Time of Incubation

As recommended by CLSI and EUCAST, antimicrobial susceptibility testing of MNPs should be performed from 16 to 20 h at 35 ± 2 °C against bacteria, and from 48 to 72 h at 25–30 °C against fungi (Table 9) [379,380]. However, most studies related to the evaluation of antimicrobial activity of MNPs have adopted an incubation period of 12–24 h at 30–37 °C for bacteria, and of 48–72 h at 37 °C for fungi (Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7). If the temperature is lowered, the time required for achieving the growth of microbes can be extended, resulting in a “false” larger diameter of inhibition (or lower MIC) [379,380]. On the other hand, high temperatures can negate the results by inhibiting the antimicrobial properties of the biofabricated MNPs [360].

With the aim of determining the bactericidal and fungicidal effects of nanoparticles as a function of incubation time, a time-kill test can be performed by incorporating a bacterial or fungal suspension and different concentrations of MNPs (e.g., 0.25 MIC, 0.5 MIC, 1 MIC, etc.) in different tubes or microplates containing the broth culture medium [319,381]. The percentage of dead cells at various time intervals (0.5 h, 1 h, 6 h, 12 h, etc.) can then be calculated relative to the growth control by determining the number of living cells present in each tube or microplate according to validated count methods [318,381]. Generally, the bactericidal effect is obtained with a lethality percentage of 90% for six hours, which is equivalent to 99.9% of lethality for 24 h. Additionally, survivor counts from collecting sequential samples are generally plotted to obtain a “time-kill curve” after an incubation period at a predetermined temperature [382,383]. Therefore, standardization of temperature and time of incubation is required to ensure the comparison of results from one study to another.

4.2.6. Size and Shape of Nanoparticles

The antibacterial and antifungal properties of MNPs depend strongly on their sizes and shapes. Small nanoparticles (diameter sizes ˂ 30 nm) show better antimicrobial activity than large nanoparticles (diameter sizes ˃ 30 nm) [384,385,386], since smaller particles offer a greater surface area for contact, favoring MNP-microbial cell interactions, as well as, more importantly, physical damage to microbial membranes. A number of nanoparticles per bacterium is required to induce cell death [387,388]. Additionally, the ability of small nanoparticles to freely permeate inside the microbial membrane is higher than that of large nanoparticles [387,388].

It has been observed that needle-shaped metal oxide nanoparticles generally exhibit significantly greater antimicrobial activity than those with a cube shape [389]. Additionally, truncated triangular silver nanoparticles were found to be more effective against E. coli than rod-shaped silver nanoparticles, while rod-shaped particles were much more effective than spherical nanoparticles [390,391].

Therefore, the size and shape of antimicrobial MNPs must be considered when comparing their MIC values and the diameter sizes of inhibition. However, a small number of authors omit to provide information relating to these physicochemical characteristics (see Table 1, Table 2, Table 3, Table 4, Table 5, Table 6, Table 7 and Table 8 and S1–S7).

The ratio between the metal salts and plant extracts used during biosynthesis must also be taken into account, since the amount of capping agents (phytoconstituents) on the surface of MNPs, as well as the size, shape and antimicrobial activities of the nanoparticles, depends on it [392,393]. The seasonal and geographical variability of plant compositions should also be considered. Hence, the collection periods and sites should be referred to in all studies, although this has not always been the case to date.

4.2.7. Zeta Potential of Nanoparticles

The zeta potential (or charge surface) is one of the critical properties of nanoparticles that can affect their stability and cell adhesion [18,394]. Balanced by oppositely charged counter ions present in the surrounding media, the surface charge of bacterial and fungal membranes is generally negative [395]. Hence, upon interaction with these membranes, green MNPs with positive surface charges may impede the growth of bacteria and fungi by preventing their attachment [385,388,396,397,398]. Moreover, the capacity of cationic nanoparticles to improve the production of reactive oxygen species, and the exertion of mechanical stress on the microbial membrane, appear to be greater than that of anionic and neutral nanoparticles [280,281].

The presence, or addition, of certain cationic agents (e.g., polymyxin, cetyltrimethyl ammonium bromide (CTAB), benzalkonium chloride and chlorhexidine digluconate) to the culture media can modify the zeta potential of bacterial and fungal cell membranes through electrostatic interactions. This may subsequently alter cell surface permeability, leading to the death of bacteria and fungi [399]. In addition, aminoglycoside antibiotics (e.g., gentamicin, neomycin, paromomycin) that are positively charged at physiological pH can synergize with all kinds of MNPs [400,401]. On the other hand, depending on their zeta potential, phytocomponents that act as stabilizing and capping agents can reduce the propensity of biosynthesized nanoparticles to aggregate, which influences their size, shape, stability and biological activity [392,402].

It should be remembered that MHB is a culture medium recommended by the US Food and Drug Administration, World Health Organization, EUCAST and CLSI for testing the susceptibility of aerobic and facultative anaerobic bacteria to antimicrobial agents [356,403]. In addition, cation-adjusted MHB 2 (CAMHB) broth, which corresponds to MHB in which divalent ions (e.g., calcium and magnesium) have been adjusted, can greatly reduce the MIC of several antibiotics (e.g., daptomycin, quinolones, aminoglycosides, and tetracycline) by affecting their mode of action and stability [356]. It is not ruled out that this phenomenon may also be observed with greenly biosynthesized MNPs, although, to the best of our knowledge, there is no study of this to date.

It is advisable to measure and indicate the zeta potential of dispersions containing MNPs and other components. As well as the effects of the interfacial potential on the antimicrobial propensity of MNPs, all the phenomena that affect the zeta potential should be studied more extensively, particularly when evaluating the efficacy of MNPs in combination with cationic antimicrobial agents.

4.2.8. pH of Culture Media

As mentioned earlier, phytoconstituents (e.g., flavonoids, tannins, alkaloids, etc.) from plant extracts used in green synthesis can cap the surface of biogenic MNPs through highly complex mechanisms of reduction of metal ions and nucleation/stabilization of reduced metal atoms [393]. All these phytochemical metabolites contribute to the antibacterial and antifungal activity of MNPs, and reduce toxicity by passivation of the surface [393,404].

Notably, if the pH of the culture medium is too low, some phytoconstituents (e.g., amino acids, proteins), and antimicrobial agents used as positive controls (e.g., quinolones, macrolides, aminoglycosides), may lose their potency, while other antibacterial drugs, such as tetracyclines, may show enhanced efficacy [405,406,407]. On the other hand, certain phytocomponents, such as caffeic, chlorogenic and gallic acids, as well as antimicrobial proteins, become unstable at high pH values, thereby losing their biological and pharmacological activity [408]. Additionally, the pH of the culture media can impinge on or modify the surface charge density and distribution of both metals and phytoconstituents capped on the surface of MNPs. This phenomenon could explain why green-synthesized silver nanoparticles and zinc nanoparticles become less active in acidic and basic media, respectively [409,410,411,412,413,414].

Therefore, the pH (and ionic strength) of culture media should be standardized to maintain the physical and biological properties of the metallic and phytochemical components of biogenic MNPs. The pH values of the culture media should range between 6.8 and 7.4, irrespective of the temperature [359].

4.2.9. Antibiotic and Antifungal Reference Standards

To assess the antimicrobial activity of MNPs, a good and common practice consists of testing them side-by-side with well-known antibiotic and antifungal references, and comparing the resultant antimicrobial activity [301,415,416]. Biosynthesized MNPs would be considered more active than a standard antimicrobial agent if the latter has a higher MIC (or lower diameter of inhibition zone) than the nanoparticles.

Depending on the MICs and the diameters of the inhibition zone, microorganisms can be considered “sensitive” or “resistant”. These values are generally compared with those mentioned in local microbiology guidelines and/or the “antibiotic and antifungal disc interpretative criteria and quality control table” proposed by EUCAST and CLSI [368,379,417,418]. The susceptibility of microorganisms to MNPs improves with decreasing MICs (or increasing diameters of inhibition) [419].

To select the most suitable antimicrobial reference test and compare its in vitro activity to MNPs, investigators can use breakpoint tables or performance standards for antimicrobial susceptibility testing [358,418,420]. Nevertheless, drug purity and disc potency (i.e., amount of drug per disc) should be considered for a more in-depth comparison of activities. Indeed, these two parameters can be reduced owing to deterioration during storage, thereby decreasing the antimicrobial efficacy of the standard test [418].

It would be interesting to simultaneously evaluate MNPs, antimicrobial standards, and test microorganisms alongside a standard microorganism having known MIC or inhibition diameter values. This strategy would enable confirmation of the reliability of the experimental conditions and the obtained results [360].

Representative (nano)antimicrobial agents can be dissolved in a suitable solvent (dilution method) or impregnated in discs (diffusion method) [303,309]. However, homogenous and reproducible disc impregnation of MNPs that are manufactured in solid (powder) or semi-solid (cream or ointment) form constitutes a bottleneck that hampers the reliable correlation of the diameter of inhibition with antimicrobial activity. To remedy this problem, the agar dilution method can be used as an alternative [307,308]. For this purpose, a volume or weight of varying concentrations of metallic nanoparticle preparations can be incorporated in different wells. Nevertheless, to obtain more comparable results, it would be necessary to standardize the volume of the wells and the quantity of the tested samples.

4.2.10. Synergistic Activity of Nanoparticles with Antimicrobial Substances

In exploring alternative approaches to improving the therapeutic management of infectious diseases, the combination of MNPs with antimicrobials (e.g., amoxicillin, azithromycin, cefotaxime, cefuroxime, chloramphenicol, clindamycin, erythromycin, fosfomycin, penicillin G, vancomycin, etc.) can be effective [400,421,422,423,424]. This strategy exhibits enhanced antibacterial and antifungal effects against different types of microbes in comparison with MNPs and antimicrobial agents alone [400,421,422,423,424].

To better evaluate this approach and ensure appropriate comparison of results from one study to another, several parameters must be considered and standardized. This particularly concerns the composition of the culture media, in terms, for example, of cationic substances (e.g., CAMHB), the spectrum of activity, and the efficacy of the selected antimicrobials, as well as the concentration ratio of the latter with MNPs.

5. Methods for Testing Anti-Biofilm Activities of Green Metallic Nanoparticles

To quantify cells in biofilm-grown microbes, many direct and indirect counting methods have been developed [425]. Direct measurement methods include viable cell enumeration using plate counts, microscopic cell counts, Coulter cell counting, flow-based cell counting and fluorescence microscopy. In addition, indirect counting methods comprise microplate assays, dry mass determination, total organic carbon, total protein, ATP bioluminescence and quartz crystal microbalance, Calgary biofilm device, biofilm ring test, etc. Of all these methods, microplate plate models are the most commonly used for the evaluation and measurement of the antibiofilm activity of green MNPs [57,425,426,427,428].

Quantitative methods of biofilm characterization are often accompanied and assisted by qualitative methods (e.g., scanning electron microscopy, scanning electrochemical microscopy) for imaging the surface roughness, morphology and spatial organization of biofilms, as well as their interaction with the environment [425,427].

5.1. Microplate Assays

As stated above, microplate plate assays remain among the most frequently used methods for evaluating the antibiofilm activities of green MNPs. Microtiter plate methods are relatively inexpensive, easy to perform, rapid and reproducible. Additionally, these methods have great flexibility due to their many variations or modifications, such as the tetrazolium salt assay and the crystal violet assay [426].

Among the most widely used tools in biology for real-time evaluation of cellular viability and metabolism in vitro, tetrazolium salt assays allow direct and indirect measures of biofilm growth via UV-visible and fluorescence spectroscopy [426]. The different tetrazolium salts are metabolically converted to formazan derivative crystals which are solubilized in dimethyl sulfoxide (DMSO) before being quantified by spectrophotometry [426]. In the crystal violet assay, biofilm cells are stained with crystal violet dyes and then infused with decoloring solution (e.g., pure ethanol, acetic acid, methanol, etc.). The resulting solution is generally transferred to clean 96-well plates to assess the optical density (OD) or absorbance at 530–600 nm using a microtiter plate reader. Homogenous resolubilization of crystals and dyes according to recommended protocols enables precise measurement of biofilm production since microplate readers measure the optical density only at one point in the middle of the well [57,429,430,431,432,433,434,435,436,437,438].

5.2. Factors Influencing the Evaluation of Antibiofilm Activities of Metallic Nanoparticles

Various static and batch-growth conditions may influence biofilm formation by different species and strains of bacteria and fungi in microtiter plates. These testing conditions include the storage of microorganisms, inoculum density, culture medium, microtiter plate, cultivation of biofilm, washing, fixation and staining. All these factors are discussed below.

5.2.1. Storage Conditions

When stored by freezing at −70 °C or by lyophilization, microorganisms generally maintain their virulence properties once thawed from storage. However, this is not the case with some fastidious bacteria and fungi (e.g., Staphylococcus spp.) which can produce mixtures of phenotypes differing in their ability to form biofilms [426]. The influence of storage conditions should be considered during the evaluation of antibiofilm activities. However, most studies relating to the evaluation of the activity of MNPs against staphylococcal-based biofilms do not mention it.

5.2.2. Type of Microorganisms

Biofilm formation depends on the type of microorganisms selected for the experiment [439]. According to the literature, MNPs are mainly evaluated against Escherichia coli, Staphylococcus aureus, Pseudomonas aeruginosa and Candida albicans. Other microorganisms include Staphylococcus epidermidis, Enterococcus faecium, Enterococcus faecalis, and Klebsiella pneumoniae. For all these microorganisms, the strain numbers assigned in international culture collections should be reported (e.g., Staphylococcus epidermidis ATCC 35984 (high slime producer) vs. Staphylococcus epidermidis ATCC 1228 (non-slime producer), or Staphylococcus aureus ATCC 29213 (methicillin-sensible) vs. Staphylococcus aureus ATCC 43300 (methicillin-resistant)). For clinical or environmental isolates, all available and relevant background and ethical information should be reported [426,440].

5.2.3. Inoculum Density

Biofilm density increases with increasing initial inoculum. The inoculum size can vary between 103 and 108 CFU/mL [57,429,430,431,432,433,434,435,436,437,438]. Nevertheless, several authors consider that a microbial cell size of 1 × 108 CFU/mL is suitable for studies dealing with the evaluation of antibiofilm activities of MNPs [51,429]. As a result, the inoculum density should be carefully standardized when evaluating antibiofilm properties, as when determining the antibacterial and antifungal activities of MNPs. However, we regret to note that the inoculum density is not given in some studies, and that it is not determined with precision in other studies.

The inoculation preparation (e.g., culturing methods) can affect the ability of microorganisms to attach to a surface [441,442]. However, information concerning inoculation preparation (e.g., concentration, temperature, time, temperature, growth phase, shaking conditions, growth media, humidity, CO2) is often not reported in published papers related to MNPs.

Cell density can also affect the spreading and clustering of certain microorganisms [443]. The percentages of clusters and of total clustered cells increase linearly with the density of inoculum (but are not time-dependent). Indeed, in the presence of biofilm-associated clusters, preexisting cell clusters that can form in cell suspension may lead to false-positive results [429]. Hence, to avoid inoculation of these preexisting clusters, cell suspensions should be broken up with a syringe fitted with a 23-gauge needle and then with a vortex [443]. These issues are of great importance and must be taken into account by researchers who are interested in evaluating the antibiofilm activities of MNPs.

5.2.4. Culture Medium

The ability of bacteria and fungi to produce biofilms under in vitro conditions depends strongly on the composition of the medium, which itself varies from one supplier to another [426]. The agar media closely resemble the surfaces (catheters, prostheses, etc.) found in in vivo situations. Hence, during biofilm formation, surface-associated bacteria and fungi can adhere better to solid growth media than to liquid culture media [444,445]. Nevertheless, in the absence of a clear-cut recommendation on the medium to be used for testing bacteria-based biofilm formation, some authors use Trytic soy broth (TSB) and brain heart infusion (BHI) broth, supplemented with glucose, sodium chloride or ethanol [429,430,431,437,446,447,448,449,450].

For fungi, yeast nitrogen base (YNB), yeast peptone dextrose (YPD), Roswell Park Memorial Institute–1640 (RPMI-1640) medium, artificial saliva medium (ASM), Sabauraund dextrose broth (SDB) or phosphate-buffered saline (PBS) can also be used, with or without supplement [429,436,438,450,451,452,453,454,455,456].

Therefore, the choice of medium for biofilm cultivation is an essential element in the process aimed at standardizing the analytical methods of antibiofilm activities of MNPs. Many efforts must be made in this direction.

5.2.5. Type of Microplates

To mimic the surfaces to which microorganisms can adhere to form biofilms, both tissue and non-tissue culture microtiter plates can be used [429]. However, cell attachment and proliferation were better on surface-treated tissue culture plates than on non-tissue culture plates [457,458]. Nevertheless, tissue-culture-treated plates from different manufacturers may provide different conditions for the cultivation and proliferation of biofilms. On the other hand, flat-bottomed microtiter plates allow better biofilm quantification than U-shaped and V-shaped microtiter plates [426,457,459]. Hence, for more transposable and comparable results, all these differences should be noted and considered by research groups. All the microplate characteristics (e.g., color of plate, number of wells, material, pre-coating conditions, etc.) should also be taken into account as part of an overall standardization process for microplate assays [426].

5.2.6. Time and Temperature of Incubation

The duration and temperature of incubation are important parameters for the cultivation of biofilms; indeed, the density of biofilms is dependent on these two parameters [443,460]. Covered with a lid, the inoculated plate should be incubated aerobically for 24 ± 0.5 h at 36 ± 1 °C under static conditions [426,429]. However, some authors incubate microtiter plates for 18 h or 20–24 h, while others incubate overnight [461,462,463]. In addition, literature reports suggest that some investigators prolong their incubation time for 48 h [426,464].

All this indicates a lack of standardization in the operating conditions of incubation. This situation can generate both false-positive and false-negative results. Even more so, it can negatively impact on the possibility of comparing the results from one study to another.

5.2.7. Washing, Fixation and Staining Steps

The washing steps aim to remove non-adherent cells and unbound dye on biofilm-containing microplates after the formation of biofilms. However, excessive washing may lead to false-negative results. In contrast, false-positive results can be obtained from insufficient washing. To avoid this, three- and four-step washing protocols are advisable [429].

A variety of methodologies and techniques can be applied during the washing step. However, some can lead to false-positive results (e.g., washing robots), while others can cause disruption of biofilm layers (e.g., mechanical plate washers). As a result, washing using micropipettes followed by emptying by flicking is considered to be a simple and effective method which is applied by several investigators [429].

Colorimetric methods have also been used to quantify metabolic activity in biofilms. For this purpose, tetrazolium salts and resazurin can be used. Compared to tetrazolium salt assay, resazurin-based quantification is inexpensive and less time-consuming. Moreover, it offers a good correlation with CFU counts. Nevertheless, one of the drawbacks of the colorimetric method using resazurin is the high lower limit of quantification (˃106–107 CFU/biofilm). An alternative approach for this method is currently available, decreasing the lower limit of quantification to 103 CFU/biofilm [429].

Extensive detachment and removal of sessile bacterial cells can result from the stringency of washing [426,429]. To prevent this generating artifacts or false results, washing must be followed by a fixation step with absolute ethanol, methanol or heat fixation at 60 °C for 1 h. After this last step, the adherent biofilm layers present in microtiter plates must be stained with dyes, such as crystal violet and alcian blue. Alcian blue stains both live and dead cells, as well as other components found in the matrix of the biofilm matrix, and is, therefore, well suited to the quantification of total biofilm biomass. However, unlike alcian blue, the crystal violet dye does not color slimy material. Other limitations of crystal violet include a lack of reproducibility, its non-specific nature in poly-microbial communities, and the absence of standardized protocols. Nevertheless, crystal violet remains the most frequently used strain for biofilm quantification [425,426,427,429].

6. Conclusions and Perspective

When searching the literature on green MNPs from 2010 to date, it appears that most of the published data are related to MNP synthesis (e.g., methods, mechanisms, influence of parameters), physicochemical characterization (e.g., size, zeta potential), and the evaluation of antimicrobial and antibiofilm properties and mechanisms of action. Of the 600 articles dedicated to antimicrobial metal nanoparticles that we have presented in this review, very few were focused on factors that affect the pharmacological properties of these MNPs and the outcomes of their in vitro evaluation.

Based on this, we set out to critically examine the experimental conditions and essential factors that can affect the antimicrobial performance of MNPs, enabling pinpointing of important MNP quality attributes for the effective development of design rules in the field of biogenic nanotechnology for tackling infectious diseases.

The present review provides, firstly, an understanding of what is commonly reported in scientific articles related to antimicrobial and antibiofilm MNPs. Secondly, this review highlights the critical procedures and parameters that influence the antimicrobial and antibiofilm evaluation of MNPs.

In addition to the lack of detailed information regarding the chemical and biological materials, as well as the laboratory equipment used, insufficient knowledge and the non-standardization of experimental protocols and laboratory conditions make it difficult to produce reproducible and reliable data or to gain detailed insights into comparative studies. This situation has been made worse by the progressive limitation in the number of words in several journals.

To address the issues arising from these multifaceted problems, the elaboration and use of minimal information guidelines or reporting standards related to the evaluation of antimicrobial and antibiofilm activities would constitute an effective strategy and approach. Minimal information guidelines should provide a guide for researchers on the necessary information that a manuscript should include for specific experiments. However, the elaboration of these guidelines requires a variety of studies, a dialog among experts and a consensus by the research community. In the interim, we have presented this draft which includes the minimum operating conditions required which would facilitate reproducibility and the reliable comparison of research data. By applying a standardized approach to experimental conditions for the evaluation of antimicrobial and antibiofilm activity, researchers will push the field of biogenic nanotechnology to new frontiers towards the development of high-value antimicrobial MNPs that can attack both existing and emerging antimicrobial resistant strains.