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Nanomaterials logoLink to Nanomaterials
. 2022 May 27;12(11):1841. doi: 10.3390/nano12111841

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

Miryam M Luzala 1,, Claude K Muanga 1,, Joseph Kyana 2,, Justin B Safari 3,4, Eunice N Zola 1, Grégoire V Mbusa 5,6, Yannick B Nuapia 7, Jean-Marie I Liesse 5,6, Christian I Nkanga 1, Rui W M Krause 4,8,*, Aistė Balčiūnaitienė 9, Patrick B Memvanga 1,2,3,10,*
Editor: Zili Sideratou
PMCID: PMC9182092  PMID: 35683697

Abstract

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.

Keywords: plant-based synthesis, metallic nanoparticles, antimicrobial and antibiofilm activities, drug-susceptibility testing methods, influencing factors

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.

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.

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.

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.

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.

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 °C
24 h
pH NM
Inoculum NM
No control
E. coli ATCC 25922
P. aeruginosa ATCC 27853
S. aureus ATCC 49476
18
15
16 mm
[87]
Perilla frutescens Leaves Silver nitrate 2 mM/plant extract 10% (9:1 v/v)
50 °C
2 h
pH NM
Spherical, rhombic, triangle, and rod
25.7 nm
Diffusion
37 °C
24 h
pH NM
Inoculum NM
Streptomycin **
E. coli
B. substilis
S. aureus
14
12
10 mm
[149]
Ocimum canum Leaves Silver nitrate 1 mM/plant extract 10% (9:1 v/v)
80 °C
15 min
pH NM
Spherical
6.1–32.1 nm
Diffusion
28 °C
24 h
pH NM
Inoculum NM
No 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 °C
24 h
pH NM
Inoculum NM
No control
B. cereus MTCC 1272
E. coli MTCC 1687
K. pneumoniae MTCC 530
Proteus mirabilis MTCC 425
P. aeruginosa MTCC 1688
S. typhi MTCC 531
S. aureus MTCC 96
12
13
14
15
11
12
11 mm
[139]
Diospyros malabarica Fruits Silver nitrate 1 mM/plant extract 20% (9:1 v/v)
25 °C
1 h
pH NM
Spherical
17.4 nm
Diffusion
37 °C
24 h
pH NM
Inoculum NM
Streptomycin 10 µg
Tetracycline 30 µg
Chloramphenicol 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 NM
pH NM
Spherical
3–25 nm
Dilution
37 °C
24 h
pH NM
Inoculum NM
No 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 °C
1 h
pH 7.2
Spherical
30 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
No control
S. typhi
Enterococcus faecalis
Shigella flexneri
Citrobacter 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 °C
24–48 h
pH NM
Irregular
26.5 nm
Diffusion
35 °C
18 h
pH NM
Inoculum NM
No control
S. aureus
P. aeruginosa
E. coli
K. pneumoniae
30
12
14
18 mm
[154]
Melissa officinalis Leaves Silver nitrate 5 mM/plant extract 25% (1:2 v/v)
25 °C
1 h
pH NM
Spherical
12 nm
Diffusion
36 °C
24 h
pH NM
Inoculum NM
No control
E. coli
S. aureus
12
13 mm
[155]
Piper betle Leaves Silver nitrate 1 mM/plant extract 10% (10:1 v/v)
Room temperature
24 h
pH NM
Spherical
Size NM
Diffusion
30 °C
24 h
pH NM
Inoculum NM
No control
B. subtilis
Klebsiella planticola
14
13 mm
[156]
Rosa canina Fruits Silver nitrate 1 mM/plant extract% NM (5:1 v/v)
85 °C
Incubation time NM
pH NM
Spherical
13–21 nm
Dilution
37 °C
24 h
pH NM
2.4 × 107 CFU/mL
No control
Bacillus cereus
E. coli ATCC 10536
S. aureus ATCC 6538
P. aeruginosa ATCC 9027
Enterococcus hirae ATCC 10541
Legionella pneumophila ATCC 33152
32
256
256
128
256
16 µg/mL
[157]
Dilution
25 °C
48 h
pH NM
2.4 × 107 CFU/mL
No control
C. albicans 128 μg/mL
Fagonia indica Callus Silver nitrate 4 mM/plant extract 2% (1:1 v/v)
20 °C
3 h
pH NM
Cubic
Size NM
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
Ciprofloxacin **
E. coli ATCC 23716
S. typhi ATCC 35664
Shigella sonnei ATCC 29930
Citrobacteramalonaticus ATCC 25405
12
13
13
12 mm
[158]
Barleria longiflora Leaves Silver nitrate 1 mM/plant extract 20% (9:1 v/v)
Temperature NM
Incubation time NM
pH NM
Spherical
2.4 ± 0.5 nm
Diffusion
37 °C
24 h
pH NM
Inoculum NM
Chloramphenicol **
Enterococcus spp.
Streptococcus spp.
Bacillus megaterium
Pseudomonas putida
P. aeruginosa
S. aureus
18
16
15
17
18
14.5 mm
[159]
Ipomoea batatas Outer peels Silver nitrate 1 mM/plant extract 40% (10:1 v/v)
55 °C
24 h
pH NM
Shape NM
Size NM
Diffusion
Temperature NM
Incubation time NM
pH NM
Inoculum NM
No control
Enterococcus feacium DB 01
S. enteritica KCCM 11806
Listeria monocytogenes ATCC 19111
B. cereus KCTC 3624
S. aureus ATCC 13565
10
11
11
11
0 mm
[160]
Oedera genistifolia Leaves Silver nitrate 0.1 mM/plant extract 20% (9:1 v/v)
Room temperature
1 h
pH NM
Spherical
10–60 nm
Dilution
37 °C
24 h
pH NM
1 × 108 CFU/mL
Ciprofloxacin **
Enterobacter cloacae ATCC 13047
Listeria ivanovic ATCC 19119
Streptococcus uberis ATCC 700407
S. aureus ATCC 29213
Vibrio spp.
Mycobacterium smergatis ATCC 19420
0.5
1
0.5
0.5
0.25
0.25 mg/mL
[161]
Derris trifoliate Seeds Silver nitrate 1 mM/plant extract 20% (20:1 v/v)
Temperature NM
Incubation time NM
pH NM
Spherical
16 ± 5 nm
Diffusion
NM
24 h
pH NM
Inoculum NM
No control
E. coli MTCC 723
K. pneumoniae MTCC 109
P. aeruginosa MTCC 424
S. aureus MTCC 96
19.5
20
36
0 mm
[162]
Ficus krishnae Stem bark Silver nitrate 1 mM/plant extract 5% (1:1 v/v)
37 °C
24 h
pH NM
Spherical
160–260 nm
Diffusion
37 °C
24 h
pH NM
Inoculum NM
No control
E. coli MTCC 45
S. typhimurium MTCC 98
S. aureus ATCC 29122
18
13
12 mm
[163]
Psidium guajava Leaves Silver nitrate 10 mM/plant extract 2% (10:1 v/v)
70 °C
1 h
pH NM
Spherical
96 ± 4 nm
Diffusion
37 ± 2 °C
48 h
pH NM
1–2 × 105 CFU/mL
No control
C. albicans ATCC 10231 14.2 mm [164]
Citrus limon Leaves Silver nitrate 2 mM/plant extract 20% (9:1 v/v)
25 °C
1 h
pH NM
Spherical
8–15 nm
Diffusion
Temperature NM
18–24 h
pH NM
Inoculum size NM
No control
Fusarium oxysporium
Alternaria brassicicola
15
10 mm
[165]
Chaenomeles sinensis Fruits Silver nitrate 1 mM/plant extract 10% (ratio NM)
80 °C
65 min
pH NM
Spherical
5–20 nm
Diffusion
37 °C
24 h
pH NM
Inoculum NM
Neomycin **
E. coli
S. aureus
14
10 mm
[166]
Persicaria odorata Leaves Silver nitrate 1 mM/plant extract 2% (10:1 v/v)
25 °C
24 h
pH NM
Spherical
11 ± 3 nm
Dilution
37 °C
18 h
pH NM
1 × 106 CFU/mL
No control
S. epidermidis ATCC 12228
MRSA ATCC 43300
3-LR ***
6-LR
[167]
Citrus reticulata Peels Silver nitrate 1 mM/plant extract 21.8% (1:1 v/v)
Temperature NM
Incubation time NM
pH NM
Spherical
45 nm
Dilution
37 °C
48 h
pH NM
1 × 105 CFU/mL
No control
Desulfovibrio spp. 3-LR [168]
Cuccuma longa Rhizome Silver nitrate 1 mM/plant extract 6.8% (4:1 v/v)
Temperature NM
24 h
pH NM
Spherical
18 nm
Dilution
37 °C
24 h
pH NM
1 × 108–109 CFU/mL
No control
E. coli
Listeria monocytogenes
4-LR
4-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 NM
Incubation time NM
pH NM
Spherical
32–85 nm
Microtiter plate
37 °C
24 h
pH NM
1.5 × 108 CFU/mL
No control
P. aeruginosa ATCC 10662 89.6% [170]
Artemisia scoporia NM Silver nitrate 1000 mM/plant extract 10% (20:1 v/v)
Temperature NM
24 h
pH NM
Spherical
10–80 nm
Microtiter plate
37 °C
24 h
pH NM
1.5 × 108 CFU/mL
No 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 °C
40 min
pH NM
Spherical
10–20 nm
Congo red agar plate
37 °C
24–48 h
pH NM
Inoculum size NM
No control
B. substilis
P. aeruginosa
NM
NM
[172]
Malva sylvestris Leaves Silver nitrate 1 mM/plant extract 20% (10:0.4 v/v)
Temperature NM
Incubation time NM
pH NM
Spherical
10–50 nm
Dilution
37 °C
40 h
pH NM
1 × 108 CFU/mL
No control
P. aeruginosa 48
P. aeruginosa B 52
62.5
62.5 μg/mL
[173]
Cannabis sativa Stem Silver nitrate 1 mM/plant extract 10% (1:1 v/v)
Temperature NM
Incubation time NM
pH NM
Spherical
20–40 nm
Microtriter plate
37 °C
24 h
pH NM
2–5 × 106 CFU/mL
No control
P. aeruginosa PA01
E. coli UTI89
S. epidermidis
6.25
12.5
50 µg/mL
[174]
Rhodiola rosea Rhizome Silver nitrate 5 mM/plant extract 10% (2:8 v/v)
90 °C
10 min
pH NM
Spherical
15–30 nm
Dilution
37 °C
24 h
pH NM
1–2 × 106 CFU/mLNo control
P. aeruginosa
E. coli
50
100 µg/mL
[131]
Flacourtia indica Leaves Silver nitrate 1 mM/plant extract 10% (1:1 v/v)
70 °C
Incubation time NM
pH NM
Spherical
45.9–64.9 nm
Congo red
37 °C
24 h
pH NM
Inoculum size NM
No control
Acinetobacter baumannii SAB5
P. aeruginosa ETPS11
K. pneumoniae SKP7
P. mirabilis PPM8
E. coli ETEC12
80
80
80
80
80 μg/mL
[116]
Dodonaea viscosa Leaves Silver nitrate 1 mM/plant extract 10% (ratio NM)
Temperature NM
18 h
pH NM
Spherical
40–55 nm
Crystal violet assay
37 °C
24 h
pH NM
1 × 107 CFU/mL
No control
C. albicans
Candida tropicalis
Candida glabrata
80
80
80%
[175]
Piper betle Leaves Silver nitrate 1 mM/plant extract 5% (19:1 v/v)
37 °C
6 h
pH NM
Spherical
156.4 nm
Microtiter plate
18 °C
Temperature NM
pH NM
Inoculum size NM
No control
Serratia marcescens
Proteus mirabilis
71
69%
[176]
Pedalium murex Seed Silver nitrate 1 mM/plant extract 5% (49:1 v/v)
Temperature NM
20 min
pH NM
Hexagonal
20–30 nm
Microtiter plate
37 °C
24 h
pH NM
Inoculum size NM
No control
Enterococcus faecalis
S. aureus
Shigella sonnei
P. aeruginosa
64
62
50
54%
[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 °C
24 h
pH 7.2
1 × 108 CFU/mL
No control
Bacillus pumulis
Enterococcus faecalis
Proteus vulgaris
Vibrio parahaemolyticus
92
84
74
62%
[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 plate
37 °C
24 h
pH NM
1 × 107 CFU/mL
No control
P. aeruginosa
S. aureus
95
90%
[179]
Allophylus cobbe Leaves Silver nitrate 5 mM/plant extract 20% (10:1 v/v)
60 °C
6 h
pH 8
Spherical
2–10 nm
Microtitre plate
37 °C
4 h
pH NM
1 × 106 CFU/mL
No control
P. aeruginosa
Shigella flexneri
S. aureus
Streptococcus pneumoniae
90
90
60
75%
[180]
Cinnamomum aromaticum NM NM
NM
NM
NM
Spherical
15–50 nm
Dilution
Temperature NM
24 h
pH 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 °C
20 min
pH NM
Spherical
37.5 nm
Congo red agar
37 °C
24 h
pH NM
Inoculum size NM
No control
P. aeruginosa
S. aureus
45
28%
[182]
Terminalia catappa Leaves Silver nitrate 10 mM/plant extract 5% (1:1 v/v)
30 °C
20 min
pH NM
Spherical
3.5–10.1 nm
Microtiter plate
37 °C
24 h
pH NM
1 × 107 CFU/mL
No control
P. aeruginosa
S. aureus
73.7
69.6%
[183]
Microtiter plate
37 °C
24 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
Diffusion
30 °C
24 h
pH NM
Inoculum size NM
No control
B. subtilis
Klebsiella planticola
13
14 mm
[156]
Musa acuminata Flowers Chloroauric acid 1 mM/plant extract 25% (9:1 v/v)
Room temperature
30 min
pH NM
Spherical
10–16 nm
Diffusion
Temperature NM
24 h
pH NMInoculum size NM
Streptomycin 10 μg
S. aureus
Enterococcus faecalis
E. coli
S. typhi
P. aeruginosa
Proteus mirabilis
0
11
7
9
9
8 mm
[192]
Zingiber officinale Roots Chloroauric acid 1 mM/plant extract 1% (2:1 v/v)
50 °C
24 h
pH NM
Hexagonal
10–20 nm
Diffusion
37 °C
24 h
pH NM
1.5 × 108 CFU/mL
No control
S. aureus
E. coli
K. pneumoniae
14
11
17 mm
[193]
Areca catechu Nut Chloroauric acid 1 mM/plant extract 5% (10:1 v/v)
80 °C
1 h
pH NM
Spherical
14 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
S. aureus
E. coli
12
14 mm
[194]
Momordica cochinchinensis Rhizome Chloroauric acid 0.01 mM/plant extract 10% (2:1 v/v)
Room temperature
24 h
pH NM
Spherical
16 ± 2 nm
Diffusion
37 ± 1 °C
24 h
pH NM
1 × 108 CFU/mL
Streptomycin 100 µg/mL
S. aureus
E. coli
B. subtilis
P. aeruginosa
19
22
19
24 mm
[195]
Plumeria alba Flowers Chloroauric acid 1 mM/plant extract 5% (5:2 v/v)
Room temperature
4 h
pH NM
Spherical
15 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli 16 mm [196]
Coleus forskohlii Root Chloroauric acid 0.1 mM/plant extract 8% (1:1 v/v)
Room temperature
2 h
pH 13
Spherical
5 nm
Diffusion
37 °C
24–48 h
pH NM
Inoculum size NM
Tetracyclin 30 μg/mL
Proteus vulgaris
Micrococcus luteus
18
14 mm
[197]
Euphorbia wallichii Leaves Chloroauric acid 1 mM/plant extract 5% (1:10 v/v)
30 °C
24 h
pH NM
Hexagonal
8 nm
Dilution
34 °C
24 h
pH NM
Inoculum size NM
Streptomycin **
E. coli
S. aureus
Bacillus pumilus
P. aeruginosa
K. pneumonia
21
15
21
17
17 mm
[198]
Coleus aromaticus Leaves Chloroauric acid 1 mM/plant extract 30% (1:1 v/v)
100 °C
30 min
pH NM
Triangular
20 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
No control
S. epidermidis
E. coli
22
27 mm
[199]
Origanum vulgare Leaves Chloroauric acid 1 mM/plant extract 10% (10:1 v/v)
85 °C
1 min
pH NM
Spherical
52 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
No control
Salmonella enteritidis ATCC 13076
E. coli ATCC 25922
Listeria monocytogenes ATCC 13932
S. aureus ATCC 6538
C. albicans ATCC 10231
10
8
10
21
28 mm
[200]
Perilla frutescens Leaves Chloroauric acid 1 mM/plant extract 10% (1:10 v/v)
30 °C
10 min
pH NM
Triangular
50 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli
B. subtilis
S. aureus
14
10
10 mm
[201]
Parkia roxburghii Leaves Chloroauric acid 1 mM/plant extract 1% (1:1 v/v)
30 °C
12 h
pH NM
Quasi-spherical
5–25 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
S. aureus
E. coli
NM
NM
[202]
Cibotium barometz Roots Chloroauric acid 1 mM/plant extract 5% (20:1 v/v)
80 °C
Incubation time NM
pH NM
Spherical
23 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Neomycin 30 µg
E. coli ATCC 10798
S. aureus ATCC 6538
Salmonella enterica ATCC 13076
P. aeruginosa ATCC 10145
16
17
13
12 mm
[203]
Mangifera indica Seed Chloroauric acid 1 mM/plant extract 10% (6:4 v/v)
80 °C
1 h
pH NM
Spherical
50 nm
Diffusion
37 °C
24–48 h
pH NM
1 × 108 CFU/mL
No control
E. coli
S. aureus
25
25 μg/mL
[204]
Rhodiola rosea Rhizome Chloroauric acid 1 mM/plant extract 10% (10:1 v/v)
80 °C
30 min
pH NM
Spherical
13–17 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
S. aureus ATCC 29213
E. coli ATCC 25922
15
12 mm
[131]
Amomum villosum Fruit Chloroauric acid 1 mM/plant extract 10% (10:1 v/v)
100 °C
60 min
pH NM
Spherical
5–10 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Neomycin **
S. aureus
E. coli
15
15 mm
[205]
Syzygium cumini Seed Chloroauric acid 1 mM/plant extract 2% (1:2 v/v)
90 °C
1 h
pH NM
Spherical
13–30 nm
Diffusion
32 °C
24 h
pH NM1 × 104 CFU/mL
Gentamicin **
E. coli
B. subtilis
S. aureus
30
33
29 mm
[206]
Hovenia dulcis Fruit Chloroauric acid 1 mM/plant extract 2.5% (5:1 v/v)
80 °C
10 min
pH NM
Spherical and hexagonal
20 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ciprofloxacin 100 µg
E. coli
S. aureus
18
19 mm
[207]
Inonotus obliquus Leaves Chloroauric acid 1 mM/plant extract 5% (19:1 v/v)
Room temperature
30 min
pH NM
Spherical
23 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
B. subtilis
S. aureus
E. coli
12
16
14 mm
[208]
Gloriosa superba Leaves Chloroauric acid 1 mM/plant extract 10% (20:1 v/v)
50–60 °C
10 min
pH 5.2
Triangular and spherical
20 nm
Diffusion
37 °C
24 h
pH NM
1.5 × 108 CFU/mL
Ampicillin 30 µg
B. subtilis ATCC 6633
E. coli MTCC 40
6.3
5.3 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 °C
2 h
pH 12
Spherical
60–80 nm
Dilution
37 °C
24 h
pH NM
1 × 105 CFU/mL
No 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 h
pH 6
Hexagonal
60–70 nm
Diffusion
35 ± 1 °C
18 h
pH NM
5 × 105 CFU/mL
No control
S. aureus ATCC 4163
E. coli ATCC 25922
P. aeruginosa ATCC 6749
S. aureus (clinical strain)
P. aeruginosa (clinical strain)
31
31
28
31
29 mm
[213]
Pongamia pinnata Seed Zinc acetate 20 mM/plant extract 20% (4:50 v/v)
60 °C
2 h
pH 12
Spherical
30–40 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ciprofloxacin 5 μg
P. aeruginosa HQ 693274.1
Bacillus licheniformis M235407.1
Vibrio parahaemolyticus HQ 693275.1
14
17
12 mm
[214]
Plectranthus amboinicus Leaves Zinc nitrate 0.05 mM/plant extract 12% (1:5 v/v)
150 °C
6 h
pH NM
Hexagonal
20–50 nm
Diffusion
37 °C
24 h
pH 7.4
1 × 105 CFU/mL
No control
S. aureus ATCC 33591 13 mm [215]
Stevia rebaudiana Leaves Zinc acetate 100 mM/plant extract 14% (1:1 v/v)
70–80 °C
2 h
pH NM
Rectangular
10–90 nm
Dilution
37 °C
24 h
pH NM
1.5 × 108 CFU/mL
No control
S. aureus
E. coli
2
2 µg/mL
[216]
Silybum marianum Seed Zinc sulfate 1 mM/plant extract 6% (1:50 v/v)
37 °C
24 h
pH 12
Flowers
60 nm
Diffusion
37 °C
24 h
pH NM
5 × 106 CFU/mL
Cefixime **
Roxithromycin **
S. aureus ATCC 6538
K. pneumoniae ATCC 1705
B. subtilis ATCC 6633
E. coli ATCC 25922
P. aeruginosa ATCC 15442
20
17
9
10
17 mm
[217]
Linum usitatissimum Root Zinc nitrate 0.1 mM/plant extract 10% (1:10 w/v)
60 °C
3 h
pH NM
Hexagonal
35 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Amoxicillin 10 µg/mL
S. aureus ATCC 6538
E. coli ATCC 15224
K. pneumoniae ATCC 4619
14
14
12 mm
[218]
Anchusa italic Flowers Zinc acetate 100 mM/plant extract 25% (1:10 v/v)
70 °C
6 h
pH NM
Hexagonal
7.6 ± 2.0 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
No control
Bacillus megaterium
S. aureus
E. coli
Salmonella typhimurium
13.6
14.6
13
14.4 mm
[219]
Conyza canadensis Leaves Zinc nitrate 150 mM/plant extract 6% (1:2 v/v)
80 °C
20 min
pH NM
Spherical
10–50 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ciprofloxacin 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 °C
15 min
pH NM
Spherical
20–25 nm
Diffusion
35 °C
24–48 h
pH NM
1 × 105 CFU/mL
Penicillin G 2 μg
Methicillin 5 μg
Vancomycin 30 μg
Gentamicin 50 μg
Streptomycin 10 μg
Ciprofloxacin 5 μg
Azithromycin 30 μg
Clotrimoxazol 25 μg
Staphylococcus spp.
Bacillus spp.
Pseudomonas spp.
Klebsiella spp.
10
0
12
11 mm
[224]
Citrus sinensis Peel Hexachloroplatinic acid 10 mM/plant extract 10% (9:1 v/v)
80 °C
24 h
pH NM
Spherical
50 nm
Diffusion
30 ± 1 °C
24 h
pH 4
Inoculum size NM
No control
Aeromonas hydrophila 4 mm [225]
Sechium edule Fruit Platinum (II) chloride 1 mM/plant extract 12.5% (1:1 v/v)
100 ± 5 °C
12 h
pH 9
Spherical
28 nm
Diffusion
37 °C
24 h
pH NM
5 × 105 CFU/mL Ciprofloxacin 30 μg Cefprozil 30 μg
B. subtilis
E. coli
25
24 mm
[226]
Spinacia oleracea Leaves Hexachloroplatinic acid 20 mM/plant extract 75% (2:1 v/v)
100 °C
24 h
pH NM
Rod
154 nm
Diffusion
37 °C24 h
pH NM
1 × 105 CFU/mL
No control
S. typhi MTCC 098 13 mm [227]
Taraxacum laevigatum Powder Hexachloroplatinic acid 10 mM/plant extract 5% (5:1 v/v)
90 °C
10 min
pH NM
Spherical
2–7 nm
Diffusion
37 °C
24 h
pH NM
5 × 105 CFU/mL
Streptomycin **
B. subtilis
P. aeruginosa
18
15 mm
[228]
Cerbera manghas Leaves Hexachloroplatinic acid 1 mM/plant extract 2% (19:1 v/v)
25 °C
2 h
pH NM
Spherical
9–12 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
Streptomycin 0.25 mg/mL
Vibrio cholerae
S. aureus
Streptococcus pyogenes
S. typhi
E. coli
20
19
13
12
11 mm
[229]
Prunus yedoensis Gum Hexachloroplatinic acid 100 mM/plant extract 25% (5:1 v/v)
80 °C
5 h
pH NM
Spherical and oval
10–50 nm
Diffusion
37 °C
48 h
pH NM
Inoculum size NM
Nystatin **
Phytophthora capsici
Phytophthora drechsleri
Didymella bryoniae
Colletotrichum acutatum
Cladosporium fulvum
0
0
0
15
18 mm
[230]
Curcuma longa Seed Hexachloroplatinic acid 1 mM/plant extract 1% (1:1 v/v)
80 °C
2 h
pH 10
Spherical
9 nm
Dilution
37 °C
3 h
pH NM
1 × 108 CFU/mL
No control
E. coli CD-496
E. coli CD-2
E. coli CD-3
E. coli CD-19
E. coli CD-549
S. aureus CD-1578
MRSA CD 489
16
16
16
16
16
64
32 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 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 *
Moringa oleifera Peel Palladium acetate 10 mM/plant extract 10% (4:1 v/v)
80 °C
5 min
pH NM
Spherical
27 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Amoxicillin **
S. aureus
E. coli
1
1 mm
[232]
Prunus yedoensis Leaves Palladium chloride 1 mM/plant extract 25% (9:1 v/v)
80 °C
30 min
pH NM
Spherical
50–150 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Amoxicillin **
B. subtilis
P. aeruginosa
6
5 mm
[230]
Cissus quadrangularis Stem Palladium chloride 0.05 mM/plant extract 10% (1:5 v/v)
37 °C
10 min
pH NM
Spherical
12–26 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli 17 mm [233]
Camellia sinensis Leaves Palladium chloride 1 mM/plant extract 1% (1:1 v/v)
40 °C
30 min
pH NM
Spherical
6–18 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
Streptomycin **
S. epidermidis S273
E. coli E266
17
14 mm
[234]
Garcinia pedunculata Leaves Palladium acetate 1 mM/plant extract 20% (2:1 v/v)
121 °C
15 min
pH NM
Spherical
2–4 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
Cronobacter sakazakii AMD04 0.3 mm [235]
Phoenix dactylifera Leaves Palladium chloride 3 mM/plant extract 10% (5:1 v/v)
37 °C
10 min
pH NM
Spherical
2–5 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
P. aeruginosa 26 mm [236]
Arabidopsis thaliana Leaves Palladium chloride 5 mM/plant extract 1% (10:1 v/v)
80 °C
24 h
pH NM
Spherical
20–40 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
S. aureus 29 mm [237]
Acacia senegalensis Gum Tetrachloropalladic acid 1 mM/plant extract 0.2% (1:1 v/v)
100 °C
6 h
pH NM
Spherical
10 nm
Diffusion
37 °C
24 h
pH NM
0.5 × 105 CFU/mL
No control
Bacillus cereus
S. aureus
Streptococcus agalatiae
18
16
17 mm
[238]
Bauhinia variegate Bark Palladium chloride 1 mM/plant extract 10% (4:1 v/v)
60 °C
30 min
pH NM
Irregular
2–9 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
B. subtilis MTCC 441
S. aureus MTCC 737
E. coli MTCC 1687
C. albicans MTCC 183
16
6
1
7 mm
[239]
Allium cepa Bulb Palladium chloride 10 mM/plant extract 10% (1:5 v/v)
100 °C
2 h
pH NM
Spherical
19 nm
Diffusion
37 °C
24 h
pH NM
1 × 106 CFU/mL
No control
Bacillus cereus
S. aureus
Micrococcus spp.
E. coli
Klebsiella spp.
Proteus spp.
36
27
40
22
18
17 mm
[240]
Filicium decipiens Leaves Palladium chloride 1 mM/plant extract 10% (9:1 v/v)
37 °C
96 h
pH NM
Spherical
2–22 nm
Diffusion
37 °C
24 h
pH NM
1 × 105 CFU/mL
Levofloxacin **
B. subtilis
S. aureus
E. coli
P. aeruginosa
12
12
27
24 mm
[241]
Phyllanthus emblica Seed Palladium acetate 870 mM/plant extract 10% (4:1 v/v)
60 °C
3 h
pH NM
Spherical
28 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Streptomycin 50 µg/mL
B. subtilis
S. aureus
P. aeruginosa
Proteus mirabilis
8.9
8.2
7.6
4.3 mm
[242]
Eucommia ulmoides Bark Palladium chloride 10 mM/plant extract 20% (5:1 v/v)
80 °C
30 min
pH 6
Spherical 2 nm Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
S. aureus
E. coli
2529 mm [243]
Delonix regia Leaves Palladium chloride 0.5 mM/plant extract 25% (9:1 v/v)
28 °C
3 h
pH NM
Spherical
2–4 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
Streptococcus mitis 12 mm [244]
Coriandrum sativum Seed Palladium chloride 10 mM/plant extract 5% (1:5 v/v)
60 °C
3 h
pH NM
Spherical
113 nm
Diffusion
37 °C
18 h
pH NM
5 × 108 CFU/mL
Streptomycin 50 µg/mL
S. aureus
E. coli
Salmonella enterica
13
8.5
10 mm
[245]
Piper betle Leaves Palladium chloride 1 mM/plant extract 20% (1:10 v/v)
30 °C
1 h
pH NM
Spherical
4 nm
Diffusion
30 °C
48 h
pH NM
Inoculum size NM
Clotrimazol 1 mg/mL
A. niger 34 mm [246]
Rosmarinus officinalis Leaves Palladium acetate 100 mM/plant extract 10% (2:1 v/v)
37 °C
24 h
pH NM
Spherical
15 nm
Diffusion
37 °C
20 h
pH NM
1 × 106 CFU/mL
Ciprofloxacin **
S. aureus
E. coli
S. epidermidis
Micrococcus luteus
24
25
21
20 mm
[247]
Difusion
32 °C
1 week
pH NM
1 × 106 CFU/mL
Nystatin **
C. albicans
Candida parapsilolis
Candida glabrata
Candida krusei
0
0
0
0 mm
Boswellia serrata Gum Palladium chloride 1 mM/plant extract 0.5% (1:1 v/v)
121 °C
30 min
pH NM
Spherical
2–9 nm
Diffusion
37 °C
24 h
pH NM
1 × 106 CFU/mL
Gentamicin 10 µg
S. aureus ATCC 25923
P. aeruginosa ATCC 27853
2628 mm [248]
Coffea arabica Powder Palladium chloride 100 mM/plant extract 8% (1:5 v/v)
60 °C
3 h
pH NM
Spherical
20–60 nm
Diffusion
37 °C
24 h
pH NM
2 × 108 CFU/mL
Ampicillin **
Enterococcus faecalis
S. typhi
S. epidermidis
12
12
12 mm
[249]
Morus alba Fruit Palladium chloride 2 mM/plant extract 20% (5:1 v/v)
80 °C
3 h
pH NM
Spherical and non-regular
50–150 nm
Diffusion
37± °C
24 h
pH NM
1.5 × 108 CFU/mL
Amoxicillin **
Listeria monocytogenes ATCC 19115
E. coli O157:H7
26
29 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 °C
3 h
pH 12
Spherical, hexagonal and oval
12–14 nm
Diffusion
37 °C
24 h
pH NM
1 × 107 CFU/mL
No control
E. coli ESβL-336
MSSA
MRSA
20
18
16 mm
[251]
Ziziphus spina-christi Fruits Copper (II) sulfate 20 mM/plant extract 6% (1:10 v/v)
80 °C
1 h
pH NM
Spherical
9 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli
S. aureus
2
2 mm
[252]
Enicostemma axillare Leaves Copper (II) sulfate 5 mM/plant extract 10% (10:1 v/v)
50 °C
24 h
pH 7
Spherical
44 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli
P. aeruginosa
K. pneumoniae
S. aureus
Proteus vulgaris
4
6
8
11
8 mm
[253]
Phyllanthus emblica Fruits Copper (II) sulfate 20 mM/plant extract 50% (3:1 v/v)
80 °C
15 min
pH 10
Flakes
15–30 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ciprofloxacin 25 μg
E. coli
S. aureus
14
14 mm
[254]
Carica papaya Leaves Copper (II) sulfate 5 mM/plant extract 10% (9:1 v/v)
60 °C
10 min
pH NM
Rod
40 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli
S. aureus
P. aeruginosa
9
9
10 mm
[255]
Sida acuta Leaves Copper (II) sulphate 1000 mM/plant extract 4% (2:1 v/v)
100 °C
5–7 h
pH NM
Spherical
50 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli
Proteus vulgaris
15
11 mm
[256]
Prosopis cineraria Leaves Copper (I) acetate 5 mM/plant extract 10% (1:1 v/v)
Room temperature
24 h
pH NM
Hexagonal
19–32 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Cefotaxim **
Proteus vulgaris
P. aeruginosa
K. pneumoniae
E. coli
S. aureus
S. epidermidis
17
18
22
22
19
23 mm
[257]
Syzygium aromaticum Buds Copper (II) acetate 1 mM/plant extract 100% (5:1 v/v)
30 °C
15 min
pH NM
Spherical
20 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
Staphylococcus spp.
E. coli
Pseudomonas spp.
Bacillus spp.
5
6
7
8 mm
[258]
Diffusion
37 °C
72 h
pH NM
Inoculum size NM
No control
A. niger
A. flavus
Penicillium spp.
5
5
6 mm
Ruellia tuberosa Leaves Copper (II) sulfate 1 mM/plant extract 5% (1:1 v/v)
100 °C
7–8 h
pH NM
Nanorod
20–100 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Streptomycin **
S. aureus
K. pneumoniae
E. coli
13
14
18 mm
[259]
Punica granatum Peels Copper (II) sulfate 50 mM/plant extract 10% (1:1 v/v)
80 °C
10 min
pH NM
Spherical
15–20 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Streptomycin **
P. aeruginosa MTCC 424
Salmonella enterica MTCC 1253
Micrococcus luteus MTCC 1809
Enterobactera erogenes MTCC 2823
19
20
20
19 mm
[260]
Asparagus adscendens Leaves Copper (II) sulfate 1 mM/plant extract 5% (10:1 v/v)
100 °C
1 h
pH NM
Spherical
10–15 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
Ampicillin 25 µg/mL
E. coli
B. subtilis
S. typhi
K. pneumoniae
S. aureus
20
18
21
18
17 mm
[261]
Gloriosa superba Leaves Copper (II) sulfate 1 mM/plant extract 5% (4:1 v/v)
60 °C
3–4 min
pH NM
Spherical
5–10 nm
Diffusion
37 °C
24–36 h
pH NM
Inoculum size NM
Ciprofloxacin 0.5 µg/µL
Klebsiella aerogenes NCIM 2098
E. coli NCIM 5051
S. aureus NCIM 5022
Pseudomonas desmolyticum NCIM 2028
15
13
6
5 mm
[262]
Cassia auriculata Leaves Copper (II) sulfate1 mM/plant extract 5% (4:1 v/v)
Room temperature
5 h
pH NM
Clusters
38 nm
Diffusion
37 °C
24 h
pH NM
1 × 108 CFU/mL
Amoxicillin **
E. coli
P. aeruginosa
S. aureus
Proteus mirabilis
Bacillus cereus
K. pneumoniae
16
10
14
16
18
14 mm
[263]
Bersama abyssinica Leaves Copper (I) acetate 100 mM/plant extract 10% (1:1 v/v)
80 °C
2 h
pH NM
Spherical
31 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Gentamicin **
S. aureus
B. subtilis
E. coli
P. aeruginosa
12
6
14
6 mm
[264]
Datura innoxia Leaves Copper (II) sulfate 1 mM/plant extract 5% (1:1 v/v)
100 °C
1 h
pH NM
Nanoclusters
90–200 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Plantomycin *
Xanthomos oryzae pv. oryzae 24 mm [265]
Zingiber officinale Rhizome Copper (II) sulfate 5 mM/plant extract 30% (5:3 v/v)
60 °C
4 h
pH NM
Spherical
31 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ciprofloxacin **
E. coli 22 mm [266]
Vaccinium arctostaphylos Fruit Copper (II) acetate/plant extract 5% (1:20 w/v)
60 °C
24 h
pH 10
Spherical
14 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Nitrofurantoïn **
E. coli 22 mm [267]
Cissus arnotiana Leaves Copper (II) sulphate 10 mM/plant extract 1% (9:1 v/v)
60 °C
4 h
pH NM
Spherical 60–90 nm Diffusion
37 °C
18 h
pH NM
Inoculum size NM
Ampicillin **
E. coli
Streptococcus spp.
Rhizobium spp.
Klebsiella spp.
22
20.2
16.3
18.3 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 °C
30 min
pH NM
Rod
16 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
P. aeruginosa
S. aureus
24
23 mm
[269]
Acacia nilotica Pods Iron (II) sulfate 100 mM/plant extract 10% (3:2 v/v)
Room temperature
24 h
pH 6
Irregular
39 nm
Diffusion
30 °C
24 h
pH NM
Inoculum size NM
No control
E. coli
MRSA
S. typhi
S. aureus
C. albicans
17
24
23
25
25 mm
[270]
Musa ornate Flowers Iron (III) chloride 1 mM/plant extract 10% (1:1 v/v)
70–80 °C
1 h
pH NM
Spherical
20–40 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
Streptococcus agalactiae
S. aureus
Salmonella enterica
E. coli
28
32
0
0 mm
[271]
Skimmia laureola Leaves Iron (III) chloride 100 mM/plant extract 10% (1:1 v/v)
Room temperature
30 min
pH NM
Spherical
56–350 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Streptomycin 200 ppm
Ralstornia solanacearum 18 mm [272]
Lagenaria siceraria Leaves Iron (III) chloride 10 mM/plant extract 5% (1:1 v/v)
40 °C
60 min
pH NM
Cubic
30–100 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ampicillin 20 mg/mL
S. aureus
E. coli
14
17 mm
[273]
Trigonella foenum-graecum Seed Iron (II) chloride 1000 mM/plant extract 5% (1:2 v/v)
Room temperature
2 h
pH 10
Spherical
∼20 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli ATCC 11775
S. aureus ATCC 6538
22
24 mm
[148]
Dodonaea vicosa Leaves Iron (III) chloride 10 mM/plant extract 20% (2:1 v/v)
50 °C
1 h
pH NM
Spherical
50–60 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
E. coli MTCC 443
K. pneumoniae NCIM 2079
B. subtilis MTCC 441
S. aureus MTCC 4032
Pseudomonas fluorescens MTCC 121
8
10
12
14
24 mm
[274]
Couroupita guianensis Peel Iron (III) chloride 100 mM/plant extract 5% (1:1 v/v)
80 °C
30 min
pH 10
Spherical
7–80 nm
Diffusion
37 °C
24 h
pH NM
1 × 105 CFU/mL
Streptomycin 1 mg
S. aureus MTCC 96
E. coli MTCC 2939
S. typhi MTCC 3917
K. pneumoniae MTCC 530
8
15
15
12 mm
[275]
Psidium guajava Fruit Iron (III) chloride 500 mM/plant extract 4% (4:1 v/v)
100 °C
1 h
pH NM
Hexagonal
27 nm
Diffusion
37 °C
18–24 h
pH NM
Inoculum size NM
Gentamicin 10 μg
Bacillus cereus
E. coli
K. pneumoniae
S. aureus
14
17
10
14 mm
[276]
Punica granatum Peel Iron (III) chloride 150 mM/plant extract 4.6% (5:2 v/v)
20 °C
5 h
pH NM
Spherical
20–90 nm
Diffusion
30 °C
24 h
pH NM
Inoculum size NM
Streptomycin **
P. aeruginosa 22 mm [277]
Argemone mexicana Leaves Iron (III) chloride 25 mM/plant extract 3% (1:1 v/v)
45 °C
12 h
pH NM
Spherical
10–30 nm
Diffusion
37 °C
24 h
pH NM
1 × 106 CFU/mL
Streptomycin 30 µg
E. coli MTCC 443
B. subtilis MTCC 425
Proteus mirabilis MTCC 441
13
18
10 mm
[278]
Ruellia tuberosa Leaves Iron (II) sulphate 1000 mM/plant extract 5% (1:1 v/v)
80 °C
30 min
pH NM
Hexagonal 53 nm Diffusion
37 °C
24 h
pH NM
1 × 106 CFU/mL
Streptomycin **
K.pneumoniae
E. coli
S.aureus
13
16
11 mm
[279]
Leucas aspera Leaves Iron (III) chloride 5 mM/plant extract 20% (1:1 v/v)
80 °C
15 min
pH NM
Irregular rhombic
117 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Ampicillin 10 µg
E. coli
K. pneumoniae
Proteus mirabilis
Salmonella enterica
Shigella flexneri
Vibrio cholera
P. aeruginosa
Bacillus cereus
S. aureus
Listeria monocytogens
10
10
11
19
22
10
20
00
11
12 mm
[280]
Eichhornia crassipes Leaves Ferrous sulphate 100 mM/plant extract 5% (1:1 v/v)
55 °C
2 h
pH 10
Rod
10–100 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
S. aureus
Pseudomonas fluorescens
E. coli
23
23
20 mm
[281]
Sida cordifolia Whole the plant Iron nitrate 10 mM/plant extract 5% (2:1 v/v)
60 °C
5 min
pH NM
Spherical
10–22 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
No control
B. subtilis
S. aureus
E. coli
K. pneumoniae
17
15
15
19 mm
[282]
Trigonella foenum-graecum Seed Iron (III) chloride 10 mM/plant extract 0.04% (1:20 v/v)
30 °C
15 min
pH NM
Spherical
11 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Streptomycin **
E. coli
S. aureus
22
19 mm
[283]
Piliostigma thonningii Flowers Iron (II) chloride 1 mM/plant extract 20% (9:1 v/v)
80 °C
2 min
pH NM
Spherical
20–100 nm
Diffusion
37 °C
24 h
pH NM
Inoculum size NM
Gentamycin **
E. coli
S. aureus
21
20 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.

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.

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.

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 Incubation
Temperature
Incubation
Time 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.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/nano12111841/s1, Table S1. Green silver nanoparticles exhibiting antimicrobial activity; Table S2. Green gold nanoparticles exhibiting antibacterial and antifungal activities; Table S3. Antimicrobial green synthesized zinc (oxide) nanoparticles; Table S4. Green platinum nanoparticles exhibiting antimicrobial activity; Table S5. Green palladium nanoparticles with antimicrobial activity; Table S6. Green copper nanoparticles exhibiting antimicrobial activity; Table S7. Antimicrobial green-synthesized iron nanoparticles [465,466,467,468,469,470,471,472,473,474,475,476,477,478,479,480,481,482,483,484,485,486,487,488,489,490,491,492,493,494,495,496,497,498,499,500,501,502,503,504,505,506,507,508,509,510,511,512,513,514,515,516,517,518,519,520,521,522,523,524,525,526,527,528,529,530,531,532,533,534,535,536,537,538,539,540,541,542,543,544,545,546,547,548,549,550,551,552,553,554,555,556,557,558,559,560,561,562,563,564,565,566,567,568,569,570,571,572,573,574,575,576,577,578,579,580,581,582,583,584,585,586,587,588,589,590,591,592,593,594,595,596,597,598,599,600,601,602,603,604,605,606,607,608,609,610,611,612,613,614,615,616,617,618,619,620,621,622,623,624,625,626,627,628,629,630,631,632,633,634,635,636,637,638,639,640,641,642,643,644,645,646,647,648,649,650,651,652,653,654,655,656,657,658,659,660,661,662,663,664,665,666,667,668,669,670,671,672,673,674,675,676,677,678,679,680,681,682,683,684,685,686,687,688,689,690,691,692,693,694,695,696,697,698,699,700,701,702,703,704,705,706,707,708,709,710,711,712,713,714,715,716,717,718,719,720,721,722,723,724,725,726,727,728,729,730,731,732,733,734,735,736,737,738,739,740,741,742,743,744,745,746,747,748,749,750,751,752,753,754,755,756,757,758,759,760,761,762,763,764,765,766,767,768,769,770,771,772,773,774,775,776,777,778,779,780,781,782,783,784,785,786,787,788,789,790,791,792,793,794,795,796,797,798,799,800,801,802,803,804,805,806,807,808,809,810,811,812,813,814,815,816,817,818,819,820,821,822,823,824,825,826,827,828,829,830,831,832,833,834,835,836,837,838,839,840,841,842,843,844,845,846,847,848,849,850,851,852,853,854].

Author Contributions

M.M.L., C.K.M. and J.K.: formal analysis, investigation, methodology, resources, validation, visualization, writing—original draft. J.B.S.: formal analysis, resources, validation, visualization, writing—original draft. E.N.Z. and G.V.M.: formal analysis, resources, validation, visualization. Y.B.N., J.-M.I.L., C.I.N., R.W.M.K. and A.B.: investigation, methodology, writing—review and editing. P.B.M.: conceptualization, formal analysis, investigation, methodology, resources, supervision, validation, writing—original draft; writing—review and editing. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available on request from the corresponding authors.

Conflicts of Interest

The authors declare that they have no competing interest.

Funding Statement

This work was partially funded by Rhodes University (South Africa) and supported by the National Research Foundation (NRF, South Africa).

Footnotes

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

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Supplementary Materials

Data Availability Statement

The data presented in this study are available on request from the corresponding authors.


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