Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1 -- 4313 2023-07-21 11:50:07 |
2 format correction -14 word(s) 4299 2023-07-24 03:24:32 |

Video Upload Options

Do you have a full video?


Are you sure to Delete?
If you have any further questions, please contact Encyclopedia Editorial Office.
Lyagin, I.; Aslanli, A.; Domnin, M.; Stepanov, N.; Senko, O.; Maslova, O.; Efremenko, E. Metal Nanomaterials and Hydrolytic Enzyme-Based Formulations. Encyclopedia. Available online: (accessed on 23 April 2024).
Lyagin I, Aslanli A, Domnin M, Stepanov N, Senko O, Maslova O, et al. Metal Nanomaterials and Hydrolytic Enzyme-Based Formulations. Encyclopedia. Available at: Accessed April 23, 2024.
Lyagin, Ilya, Aysel Aslanli, Maksim Domnin, Nikolay Stepanov, Olga Senko, Olga Maslova, Elena Efremenko. "Metal Nanomaterials and Hydrolytic Enzyme-Based Formulations" Encyclopedia, (accessed April 23, 2024).
Lyagin, I., Aslanli, A., Domnin, M., Stepanov, N., Senko, O., Maslova, O., & Efremenko, E. (2023, July 21). Metal Nanomaterials and Hydrolytic Enzyme-Based Formulations. In Encyclopedia.
Lyagin, Ilya, et al. "Metal Nanomaterials and Hydrolytic Enzyme-Based Formulations." Encyclopedia. Web. 21 July, 2023.
Metal Nanomaterials and Hydrolytic Enzyme-Based Formulations

Combination of metals and enzymes as effective antifungal agents is currently being conducted due to the growing antifungal resistance problem. Metals are attracting special attention due to the wide variety of ligands that can be used for them, including chemically synthesized and naturally obtained variants as a result of the so-called “green synthesis”. The main mechanism of the antifungal action of metals is the triggering of the generation and accumulation of reactive oxygen species (ROS). Further action of ROS on various biomolecules is nonspecific. Various hydrolytic enzymes exhibit antifungal properties by affecting the structural elements of fungal cells (cell walls, membranes), fungal quorum sensing molecules, fungal own protective agents (mycotoxins and antibiotics), and proteins responsible for the adhesion and formation of stable, highly concentrated populations in the form of biofilms. 

green synthesis MOFs amyloid proteins prionase mycotoxins growth inhibition

1. Introduction

The accumulation of information about the role that microscopic fungi can play in the development of a number of negative processes affecting human health [1][2][3] has led to increasing interest in antifungals that can control and reduce the growth, as well as the metabolic activity, of these biological objects, especially those associated with pathogens [4]. The seriousness of these tasks is increasing due to the fact that in some cases, fungal cells may develop resistance to the chemical formulations used against them [5][6][7].
A number of current scientific studies are related to the development of effective antifungals [8]. Among the new trends in the development of effective antifungals, the prospects of a possible combination of various chemical compounds [7] with different mechanisms of action on fungal cells are being considered. This approach can enable researchers to overcome the development of adaptive processes in fungi and, possibly, reduce the doses of the substances used, increasing the effectiveness of their action in such combinations. When implementing such a combined approach to suppressing the growth and metabolic activity of fungi, the main question arises about what is better to combine with what, and what may be unpromising. One of the possible answers to this question is based on the use of metal nanomaterials such as metal-nanoparticles, metal-organic frameworks, etc., to which no resistance is formed by most microorganisms since the mechanism of suppression of biological processes is primarily associated with the generation of reactive oxygen species (ROS) in the cells. Metals such as Ag, Cu, Fe, Zn, Se, Ni, Au, Zr, Ce, Ti, and Pd have been studied in regard to compounds possessing antifungal activity [9][10][11][12]. At the same time, current scientific research on the antifungal properties of metals is mainly focused on the study of Ag and Au nanoparticles (NPs) [10][11][12][13][14][15] since the antimicrobial effectiveness of their action has been known for a long time.
Among the various organic synthetic ligands for the metals used in research in this direction, the so-called “green synthesized” metal-containing NPs should be noted. These “green synthesized” metal-containing NPs are formed inside the cells of microorganisms (bacteria, fungus, yeast) in vivo or using plant extracts, polysaccharides of phototrophic microorganisms, and extracellular enzymes of mycelial fungi [10][14][15][16][17]. “Green synthesis” is an environmentally friendly synthesis technique that avoids the formation of undesired by-products and costs less. Moreover, it was found that “green synthesis” makes it possible to obtain NPs with identical antifungal properties compared to similar chemically synthesized metal-containing compounds that are, in some cases, superior to them [17].
It is known that the combination of metal NPs with known chemical fungicides makes it possible to reduce the minimum inhibitory concentration (MIC) of the latter by more than eight times [17]. However, despite this researchers decided to consider the possibility of combining metal-containing compounds with biological molecules having catalytic properties, in particular, with various enzymes exhibiting antifungal activity instead of chemically synthesized fungicides. It has been previously shown that the efficiency of the use of metal NPs can be increased by combining them with cyclic peptides that exhibit antifungal properties [18]. Unlike peptides that exhibit antimicrobial activity, the enzymes have catalytic activity [19], which allows them not just to trigger destructive processes against fungi but to repeatedly participate in these acts of biocatalysis, deepening antifungal processes. In addition, a wide substrate range of action of the enzymes themselves allows researchers to consider the possibility of not only their destructive activity against fungal cells but also against the most important fungal molecules involved in the formation of their quorum sensing (QS) and adhesion [20] and molecules that ensure their own safety (antibiotics [21] and mycotoxins [22]).

2. Antifungal Agents Based on Metal Nanoparticles, Metal–Organic Frameworks and Their Composites

Multiple antifungal agents have been developed to date on the basis of metal nanoparticles (NPs) and/or metal–organic frameworks (MOFs) ( [11][12][23][24][25][26][27][28][29][30][31][32][33][34][35][36][37], Figure 1).
Figure 1. Some representative metal NPs and MOFs with antifungal activities. Crystal structures of Ag (1741252), ZnO (13950), Fe3O4 (1612598), HKUST-1 (2091261), MIL-53-Fe (2088536), and UiO-66 (2054314) were obtained from CCDC, then expanded in Mercury (v.4.2.0, CCDC, Cambridge, UK) and visualized in PyMOL (v.1.7.6, Schrödinger Inc., New York, NY, USA). Water-accessible molecular surface is indicated by light grey while atoms are colored by element: Ag–grey, Zn–slate, O–red, Fe–orange, C–deep blue, H–white, Zr–cyan.
Table 1. Antifungals based on metal nanoparticles (NPs), metal–organic frameworks (MOFs), and their composites *.
Antifungal Agent [Reference] Target of Action Antifungal Activity Efficiency of Antifungal Action
ZrO2-Ag2O (14–42 nm) [23] Candida albicans,
C. dubliniensis,
C. glabrata,
C. tropicalis
The growth rate inhibition 89–97% inhibition
WS2/ZnO nano-hybrids [24] C. albicans Inhibition of biofilm formation 91% inhibition
CuO@C (36–123 nm) [25] Alternaria alternata,
Fusarium oxysporum,
Penicillium digitatum,
Rhizopus oryzae
Inhibition of the hydrolytic activity of fungal enzymes used by them for their own metabolism Inhibition (100 μg/mL) of cellulases and amylases secreted by fungi: 38% and 42% for A. alternata, 39% and 45% for F. oxysporum, 24% and 67% for P. digitatum, and 20% and 24%for R. oryzae, respectively
ZnO NPs [26] C. albicans,
Aspergillus niger
Inhibition of growth Large enough zone of growth absence (8-9 mm)
ZnO NPs (20–45 nm) [27] Erythricium salmonicolor Notable thinning of the hyphae and cell walls, liquefaction of the cytoplasmic content with decrease in presence of a number of vacuoles Significant inhibition (9–12 mmol/L) of cell growth
ZnO–TiO2 NPs (8–33 nm) [28] A. flavus High level of ROS production and oxidative stress induction. Treated objects have a lower count of spores and damaged tubular filaments and noticeably thinner hyphae compared to the untreated fungi Fungicidal inhibition (150 μg/mL) zone is 100 %
ZnO NPs (40–50 nm) [29] C. albicans High level of ROS production MIC = 32–64 μg/mL
MFC = 128–512 mg/mL
Fe2O3 NPs (10–30 nm) [30] Trichothecium roseum,
Cladosporium herbarum,
P. chrysogenum,
A. alternata,
A. niger
Inhibition of spore germination MIC = 0.063–0.016 mg/mL
Fe3O4 NPs (70 nm) [31] C. albicans Inhibition of cell growth and biofilm formation MIC = 100 ppm
MFC = 200 ppm
Cu-BTC (10–20 µm) [32] C. albicans,
A. niger,
A. oryzae,
F. oxysporum
ROS producing, the damage of the cell membrane Inhibition of C. albicans colonies is 96% by 300 ppm and up to 100% by 500 ppm. Inhibition growth of F. oxysporum and A. oryzae is 30% with 500 ppm. No significant effect on the A. niger growth.
HKUST-1 or HKUST-1 NPs (doped with NPs of Cu(I)) (49–51 nm) [33] A. niger, F. solani,
P. chrysogenum
Appearance of Cu+2 inhibiting of cell growth 100% growth inhibition of F. solani by 750–1000 ppm and P. chrysogenum by 1000 ppm; for A. niger—no inhibition
[Cu2(Glt)2(LIGAND)] (H2O) [34] C. albicans,
A. niger spores
The apoptosis-like fungal cell death, ROS production 50–70% death of C. albicans and 50–80% germination inhibition of A. niger at 2 mg/mL of the MOFs
MIL-53(Fe) and Ag@MIL-53(Fe) composite [35] A. flavus Inhibition of cell growth MIC = 40 μg/mL for the MIL-53(Fe);
MIC = 15 μg/mL for the Ag@MIL-53(Fe)
MOF on the basis of Ce and 4,4′,4″-nitrilotribenzoic acid [11] A. flavus,
A. niger,
Aspergillus terreus,
C. albicans,
Rhodotorula glutinis
Enzyme-like activity: catalase, superoxide dismutase, and peroxidase Inhibition efficiency of 93.3–99.3% based on the colony-forming unit method
TiO2 co-doped with nitrogen and fluorine (200–300 nm) [12] F. oxysporum Peroxidase-like activity, production of ROS under light irradiation 100% inhibition of fungal growth
Fe3O4@MoS2-Ag (~428.9 nm) [36] C. albicans Peroxidase-like activity 80% damage of cell membranes
CoZnO/MoS2 nanocomposite [37] A. flavus Peroxidase-like activity under light irradiation MIC = 1.8 mg/mL
* BTC—1,3,5-benzenetricarboxylate; Glt—glutarate; HKUST-1—type of MOFs composed of [Cu3(BTC)2(H2O)3]n; MFC—minimum fungicidal concentrations; MIC—minimum inhibitory concentration; MIL-53(Fe)—type of MOFs composed of [Fe4(OH)(1,4-benzenedicarboxylate)4]; LIGAND—1,2-bis(4-pyridyl)ethane, 1,2-bis(4-pyridyl)ethylene, or 1,3-bis(4-pyridyl) propane.

3. Enzymes as Antifungal Agents

3.1. Antifungal Enzymes Using Cell Structural Components of Fungi as Substrates

Discussing the possibilities and prospects for the use of various enzymes in the composition of antifungals, it should be noted that their diversity is determined by the spectrum of targets on which these enzymes can have notable effect, leading to a halt in the fungal growth, disruption of metabolism and death of fungi. Such targets for enzymatic action may include: structural elements of fungal cells (cell walls, membranes) [38], nucleic acids [39], fungal Quorum Sensing molecules (QSMs) regulating fungal resistance to various negative factors and protect them (mycotoxins, antibiotics) [40]), peptides and proteins involved in the formation of stable forms such as biofilms (adhesives, hydrophobins) (Figure 2).
Figure 2. Enzymes (pink—chitinase (PDB ID: 1edq); yellow—keratinase (PDB ID: 5wsl); green—lactonase AidC (PDB ID: 4zo2); blue—peroxidase (PDB ID: 1mnp) with antifungal activities due to their catalytic action on different targets as substrates.
Enzymes are interesting as antifungal agents because they are proteins present in various natural sources (plants, microorganisms, animal tissues), which are designed to protect living objects from the effects of fungi. The recombinant forms of necessary enzymes can be produced in various host cells.
The analysis of enzymes exhibiting a variety of antifungal activities indicated that most of them were hydrolases acting on polysaccharides present in the structure of the fungal cell wall or involved in the formation of biofilms. The greatest effect was observed in the case of chitinases [41][42][43][44][45][46][47][48][49][50][51][52], among which there were both exo- and endochitinases. At the same time, their simultaneous presence in the enzyme complexes used to suppress the growth of fungi was the most successful [51][53].
As a number of studies have shown [53][54][55], such a combination of chitinases with different substrate ranges can be successfully supplemented by the action of other hydrolytic enzymes (proteases and glucanases) [54][55][56][57][58], which use molecules performing the role of structural elements of the fungal cell wall and membranes as substrates. However, it should be emphasized that not all chitinases known today [59] can be used as antifungal agents since the diverse structure of fungal polysaccharides is characterized by the presence of various glycoside residues of different lengths and often does not correspond to the preferences of those substrate specificities possessed by most of these enzymes. In addition, the levels of biosynthesis of these enzymes cannot meet the needs that arise even when studying their properties. In this regard, it becomes necessary to resort to obtaining their recombinant forms, where the most commonly used cells for this purpose are E. coli BL21 (DE3) [43][44][50][60][61].
It is important to note that yeast cells are usually difficult to destroy since their cell walls can form capsules or resistant spores. Using a complex of lysing enzymes such as Lyticase (a mixture of β-(1-3)-glucan-laminar-ipentaohydrolase, β-(1-3)-glucanase, protease, and mannanase), DNA can be extracted from yeast cells [55]. The activity of this complex induces the partial formation of spheroplasts; subsequently, the spheroplasts are lysed with the release of DNA.
Nucleases hydrolyzing the RNA and DNA of fungi attract particular attention among enzymes that have antifungal activity [62][63][64]. The use of several nucleases at once [54][55][58] or nuclease in combination with glucanase [64] leads to the fact that not only is the growth of fungal cells stopped, but membrane destruction (permeabilization and depolymerization) is observed. This reduces the membrane potential of mitochondria, leading to the degradation of target cell nucleic acids and the death of microbial cells. When giving priority to hydrolases when necessary to influence fungi, it can be emphasized that they generally have the potential for antifungal effects in a fairly wide range of pH values (3.0–11.5) and temperatures (up to 80 °C). At the same time, it should be noted that the activity of hydrolases strongly depends on the presence of various metals in the media of their functioning [43][44][47][48][52][57][65]. In such media, the most attractive options are those combinations of enzymes and metals that can significantly increase the effectiveness of the antifungal action of hydrolases. Among the metal ions, which in the largest number of studies have had a stimulating effect on the activity of hydrolases, Cu2+ [47][48] and Ca2+ [52][57][60][61] should be singled out, although their positive effect is not at all unambiguous, and in some cases, they had the opposite (inhibitory) effect on the hydrolytic activity of enzymes. At the same time, the positive results obtained during enzymatic reactions directed against fungi in environments in the presence of metals indicate the expediency of searching for possible combinations of metals and enzymes in the development of new antifungal formulations.
Oxidoreductases, in particular, peroxidases are standing in second place after hydrolases in popularity among enzymes used as potential antifungal agents [66][67]. These enzymes catalyze the oxidation of fungal molecules by reducing hydrogen peroxide (H2O2). The limitations in the use of these enzymes as antifungal agents are associated with a lower efficiency of their action in comparison with hydrolases and the need to introduce additional H2O2 into the medium with fungi.

3.2. Enzymes Hydrolyzing Fungal Proteins with Amyloid Characteristics

Special attention should be paid to the fact that yeast and mycelial fungi are able to form amyloids, which are unbranched fibrils consisting of monomers stacked on top of each other and stabilized by intermolecular β-layers. For example, monomers of hydrophobins of class I, small surface-active proteins produced by fungi, form amyloid fibrils that perform many functions [68]. It is known that the specific functions of hydrophobins synthesized by fungi can enhance their pathogenicity. Thus, A. fumigatus can cause invasive aspergillosis in patients with weakened immunity due to the amyloid-forming ability of hydrophobin RodA [69][70]. The formation of amyloid by hydrophobin MPG1 in M. oryzae contributes to rice pyriculariosis [71]. One of the most well-described examples of amyloid proteins in yeast cells is the Cdc19 protein from S. cerevisiae, which, in the absence of glucose, self-aggregates into an amyloid-like aggregate to avoid degradation under stressful conditions [72].
It is known that the yeast cells of C. albicans, often used in studies of antifungals, also contain proteins with amyloid characteristics. Thus, the proteins As1, As3, and As5 from the ALS-type adhesion family have the ability to self-aggregation. The presence of an amyloid sequence in the monomers of these proteins leads to the formation of hydrophobic nanodomains that promote the cell adhesion of C. albicans on biotic or abiotic surfaces and improve their ability to form biofilms [73][74]. It is assumed that Sap6, Rbt1, Page59, and Pga62 proteins, as well as adhesins, play a significant role in the appearance of C. albicans biofilms due to the presence of an amyloid-forming sequence in their structures [75][76][77][78].
Today, due to their ability to be transmitted from “mother” cells to “daughter” cells, yeast prions are classified as infectious, for example [URE3] and [PSI+], HET and HELLP in S. cerevisiae, Podospora anserina, and Chaetomium globosum cells, correspondently [79]. The presence of similar conditions for the formation of yeast prions and common molecular properties with pathogenic human amyloids has now led to the creation of models of neurodegenerative diseases based on yeast prions. The methods of their regulation are being investigated in order to develop new effective therapeutic agents and approaches to the treatment of diseases associated with prion proteins [80]. In this regard, the interest in enzymes capable of hydrolyzing amyloid aggregates formed by fungi is huge. This is due to the possible development of antifungals that reduce the level of biofilm formation and the potential use of enzyme-containing formulations for the treatment of neurodegenerative diseases in humans. Information about such proteases hydrolyzing amyloid proteins is presented in [81][82][83][84][85][86][87][88][89][90][91][92][93][94][95][96].
Discussing the prospects for the possible use of enzymes hydrolyzing fungal amyloid proteins, it should be noted that so far there are a few such studies. The ability of several proteolytic enzymes, such as subtilisin, keratinases, and proteinase K, to degrade yeast prion aggregates of protein Sup35NM under various conditions was investigated [91][92][93][94]. It has been shown that hexameric AAA+-ATPase (Hsp104), which is a yeast chaperone, is involved in the fragmentation of large fungal amyloid fibrils. It is believed that the direct binding of Hsp104 to amyloid fibrils prevents the reproduction of yeast prions. Since Hsp104 is absent in the cells of multicellular animals, including mammals, the possibility of constructing variants of Hsp104 with the potential for use for the degradation of abnormal human proteins is being investigated [80].
Despite the limited number of studies in the field of enzymatic degradation of yeast prions, a number of proteolytic enzymes are known today that can degrade prion proteins and amyloids associated with human diseases, including subtilisin-like serine proteases TK-SP from hyperthermophilic archaeon T. kodakarensis [81], nattokinase from Bacillus subtilis natto [82], subtilisin 309 and protease from B. lentus [83][85], two prionzymes from B. subtilis and B. lentus [84][86], subtilisin-like protease MSK103 from B. licheniforms [87], enzyme E77 from Streptomyces sp. [88], subtilisin-homolog pernisine from the extremophile archaea Aeropyrum pernix [89], and serine protease from lichens [90].
Multiple metalloenzymes have been reported to have an important role in the degradation of Aβ [95][96], including two metal-activated keratinases, Ker1 and Ker2, from an actinomycete Amycolatopsis sp. MBRL 40; NEP—a zinc-dependent metalloprotease, cleaving various vasoactive peptides; and IDE—another zinc-dependent metallopeptidase, which could cleave insulin and amyloid Aβ. The ability to cleave amyloid precursor proteins has been confirmed in Zn-containing transmembrane metalloproteases [96]. At the same time, the influence of redox-active metals such as Cu and Fe (affecting the pathogenesis of Alzheimer’s disease) was established, which consists in increasing the biosynthesis of the metalloproteases under discussion. The influence of the same metals on the activity of these enzymes has not yet been investigated but is of great scientific and practical interest.
Despite the fact that there is still no effective enzymatic formulation for the cleavage of prion proteins, new proteolytic enzymes continue to be discovered and studied, the prionase activity of which still needs to be investigated [97][98][99]. Researchers focus readers’ attention on such enzymes as a potential basis for the development of new antifungals, probably with some anti-neurodegenerative effect.

3.3. Enzymes Hydrolyzing Mycotoxins, Antibiotics, and QS Molecules (QSMs) of Fungi

To date, a significant amount of information has been accumulated about QS in the cells of various fungi and molecules that are produced by the fungi themselves in response to an increase in their concentration per unit volume. These QSMs are produced in order to trigger the processes of fungal cell transition to a state of stable intercellular communication, synchronization of the functions of multicellular populations, and biochemical changes in the cells themselves [100][101][102][103][104]. The ability of individual enzymes to catalyze the hydrolysis of fungal QSMs allows them to be attributed to the so-called Quorum Quenching enzymes (QQE). Gluconolactonase- [105]) and hexahistidine-containing organophosphorus hydrolase (His6-OPH) [106][107] esterases [108][109]) have been identified as such enzymes acting against fungi today.
Discussing the potential of these enzymes as candidates for inclusion in combined antifungals with metal-containing compounds, it can be noted that for His6-OPH, such possibilities have already been demonstrated and proved promising, while Ta NPs [110][111] appeared to be the most effective option for such a combination. However, so far, such combined antimicrobials have been investigated only against bacterial cells [112], and their effectiveness against fungal cells has yet to be confirmed.
Interesting use cases for combining with metal-containing compounds are enzymes that carry out the destruction of mycotoxins synthesized by fungi in the CFR state. At the same time, it should be noted that, as in the case of CSM hydrolysis, among the enzymes that carry out the destruction of various mycotoxins (zearalenone, patulin, deoxynivalenol, ochratoxin), there are all the same enzymes, namely lactonases, esterases, lipases [22], and His6-OPH [39][113]. In this regard, with their involvement in combined antifungal formulations, a very interesting option may turn out to provide a multi-targeted action due to the promiscuous activities of these enzymes.
Continuing to analyze possible variants of enzymes that can be considered as candidates for creating combined variants with metal NPs, it is undoubtedly necessary to pay attention to enzymes that are able to catalyze the hydrolysis of antibiotics synthesized by fungi among other secondary metabolites in their QS state. Here, the undisputed leaders are β-lactamases, known to everyone due to studies of bacterial antibiotic resistance to natural and semi-synthetic penicillins and cephalosporins [114].
It is interesting to note that QQE including His6-OPH are close “relatives” for metallo-β-lactamases [115]. Moreover, the structural analogy revealed between phosphotriesterase (of the same His6-OPH) and some nucleases indicate that all these enzymes can catalyze to one degree or another similar reactions with a certain preference for individual substrates. Since these enzymes have been mentioned here more than once in connection with their various targets of action in fungal cells, their use in research on the development of new antifungals may be not only new but also promising. Surprisingly, an active search for data on the use of metallo-β-lactamases in the content of any antifungals to give them a number of catalytic activities, as discussed above, did not reveal any.
It should be noted here that many of mentioned enzymes contain different transition metals, particularly Zn(II), Mn(II), and Fe(II)/Fe(III) in their active sites [115], which can be positively taken into account when creating combinations with metal-containing compounds since there are fungi sensitive to these metals. In addition, the combination of these enzymes with metal-containing compounds that are not embedded in the active site of enzymes but can exhibit significant antimicrobial activity at low MIC values [110][111] looks interesting and promising.

4. Combination of Antifungal Enzymes and Metal-Nanoparticles

It is known currently that many sources and types of enzymes can be used to prepare antifungal formulations with metal NPs: bacterial keratinase [116] and chitinase [117]; archaeal protease and lipase [118]; fungal β-1,3-glucanase, N-acetylglucosaminidase, chitinase, and acid protease [119][120], etc. Such formulations can possess secondary antioxidant [116][117] and/or specific inhibitory activity [116]. The additional antibacterial action mode of these combinations is widely present [116][118][120][121] .
“Green synthesized” metal NPs are of great interest for the production of enzyme formulations [116][118][119][120]. β-1,3-glucanase(s) and, to a lesser extent, N-acetylglucosaminidase(s) are prevalently adsorbed by Ag NPs as compared to chitinase(s) and acid protease(s) [119]. Altogether, these enzymes on Ag NPs not only inhibit mycelium growth but also prevent the formation of sclerotia thereby leading to lifecycle arrest.
Interestingly, the “un-capping” of Ag NPs (i.e., desorption of enzymes) leads to a detectable increase of their size and is likely to be a result of their aggregation [120]. At the same time, the negative net charge of “uncapped” Ag NPs argues for the substitution of enzymes for sodium dodecylsulfate used as a solubilizer. This can contribute to the increased toxicity of such “un-capped” NPs towards non-target organisms and cell lines [120]. Surprisingly, “un-capped” Ag NPs are ineffective in a mycelium growth test and only decrease the number of sclerotia by twofold as compared to the control experiment without any effector.
Similar to germination, the formation of sclerotia is known to be regulated by multiple genes though there are a lot of gaps about this process [122]. As a result, the biochemical composition of the cell wall changes dramatically; for example, the most abundant components of Sclerotium rolfsii hyphae—polysaccharides and lipids—shift by 1.5–2 times (down and up, respectively), while unhydrolyzable compounds (so-called ‘melanin-like pigments’) increase numerously and become the second prevalent subclass (after polysaccharides). The last ones have been shown to propagate resistance of sclerotia towards environmental factors and, for example, to slaughter via the hydrolytic action of extracellular glucanases and chitinases [123]. Moreover, the leakless thick rind can be formed from such melanized cells on the sclerotia surface [124], further limiting enzymatic hydrolysis and antifungal penetration. Thus, polyphenol-degrading activity may be useful in addition to antifungal formulation. Another rational functionality in such formulation(s) to treat sclerotia appears to be the antioxidant activity discussed previously since ROS also affects sclerotial development somehow [122].
During field trials of chitinase-based formulation against filamentous fungi [117], it was found to be slightly less effective than the same formulation with a live biocontrol agent (Streptomyces cellulosae). This may be a consequence of differing profiles of protective gene modulation in the plant by these formulations.
As was determined for peptide melittin, a slow release of active compound from the Zn-MOF matrix occurs and the maximal amount (60%) is released at pH 6 during 24 h [125]. The antifungal activity of melittin is naturally decreased threefold during its encapsulation within Zn-MOF at 30 wt.% loading. However, lactoferrin added to such a formulation improves it almost twofold. Altogether, yeast adhesion to the surface during biofilm formation and (pseudo)hyphal transformation are inhibited.
Melittin is known to disturb membranes of different (micro)organisms, activate several transmembrane receptors, depolarize membranes, etc. Some of these effects are also manifested in the composite formulation [125]. Moreover, lactoferrin being a transporter of iron ions and having other possible activities [126] greatly improves the antifungal activity of melittin, especially towards pre-formed biofilms [125]. The synergic action of lactoferrin and melittin can also be detected using an animal infection model in vivo. Lactoferrin can bind to the fungal cell surface itself and affect biofilm formation and yeast-to-hyphal transition in combination with conventional drugs [127]. Thus, lactoferrin and melittin may interact with multiple and differing targets on the yeast cell wall and within the cell while amplifying the antifungal activity of each other.
Some toxicity was shown for Ag NPs toward the lung fibroblasts of Chinese hamsters, the embryo fibroblasts of albino Swiss mice, human aneuploid immortal keratinocytes, and the roots of onions [119][120]. Moreover, such formulations affect the soil microbial (bacteria and fungus) community in situ after single exposure for at least 360 days [120]. It is interesting that the toxicity of such polypeptide as melittin toward the macrophage cell line from a mouse tumor is greatly decreased within Zn-MOF formulation [125]. However, since doses of melittin in free and encapsulated form were discrepant then, total removal of toxicity cannot be concluded now.


  1. Fisher, M.C.; Gurr, S.J.; Cuomo, C.A.; Blehert, D.S.; Jin, H.; Stukenbrock, E.H.; Stajich, J.E.; Regine Kahmann, R.; Boone, C.; Denning, D.W.; et al. Threats posed by the fungal kingdom to humans, wildlife, and agriculture. MBio 2020, 11, e00449-20.
  2. Garg, D.; Muthu, V.; Sehgal, I.S.; Ramachandran, R.; Kaur, H.; Bhalla, A.; Puri, G.D.; Chakrabarti, A.; Agarwal, R. Coronavirus disease (COVID-19) associated mucormycosis (CAM): Case report and systematic review of literature. Mycopathologia 2021, 186, 289–298.
  3. Raut, A.; Huy, N.T. Rising incidence of mucormycosis in patients with COVID-19: Another challenge for India amidst the second wave? Lancet Respir. Med. 2021, 9, e77.
  4. World Health Organization. WHO Fungal Priority Pathogens List to Guide Research, Development and Public Health Action; World Health Organization: Geneva, Switzerland, 2022; p. 48. Available online: (accessed on 29 May 2023).
  5. Robbins, N.; Caplan, T.; Cowen, L.E. Molecular evolution of antifungal drug resistance. Annu. Rev. Microbiol. 2017, 71, 753–775.
  6. Fisher, M.C.; Hawkins, N.J.; Sanglard, D.; Gurr, S.J. Worldwide emergence of resistance to antifungal drugs challenges human health and food security. Science 2018, 360, 739–742.
  7. Rabaan, A.A.; Sulaiman, T.; Al-Ahmed, S.H.; Buhaliqah, Z.A.; Buhaliqah, A.A.; AlYuosof, B.; Alfaresi, M.; Al Fares, M.A.; Alwarthan, S.; Alkathlan, M.S.; et al. Potential strategies to control the risk of antifungal resistance in humans: A comprehensive review. Antibiotics 2023, 12, 608.
  8. WHO. WHO Releases First-Ever List of HEALTH-Threatening Fungi; World Health Organization: Geneva, Switzerland, 2022; Available online: (accessed on 30 May 2023).
  9. Cruz-Luna, A.R.; Cruz-Martínez, H.; Vásquez-López, A.; Medina, D.I. Metal nanoparticles as novel antifungal agents for sustainable agriculture: Current advances and future directions. J. Fungi 2021, 7, 1033.
  10. Dananjaya, S.H.S.; Thao, N.T.; Wijerathna, H.M.S.M.; Lee, J.; Edussuriya, M.; Choi, D.; Kumar, R.S. In vitro and in vivo anticandidal efficacy of green synthesized gold nanoparticles using Spirulina maxima polysaccharide. Process Biochem. 2020, 92, 138–148.
  11. Abdelhamid, H.N.; Mahmoud, G.A.E.; Sharmouk, W. A cerium-based MOFzyme with multi-enzyme-like activity for the disruption and inhibition of fungal recolonization. J. Mater. Chem. B 2020, 8, 7548–7556.
  12. Mukherjee, K.; Acharya, K.; Biswas, A.; Jana, N.R. TiO2 nanoparticles co-doped with nitrogen and fluorine as visible-light-activated antifungal agents. ACS Appl. Nano Mater. 2020, 3, 2016–2025.
  13. Wen, H.; Shi, H.; Jiang, N.; Qiu, J.; Lin, F.; Kou, Y. Antifungal mechanisms of silver nanoparticles on mycotoxin producing rice false smut fungus. Iscience 2023, 26, 105763.
  14. Malik, M.A.; Batterjee, M.G.; Kamli, M.R.; Alzahrani, K.A.; Danish, E.Y.; Nabi, A. Polyphenol-capped biogenic synthesis of noble metallic silver nanoparticles for antifungal activity against Candida auris. J. Fungi 2022, 8, 639.
  15. Soleimani, P.; Mehrvar, A.; Michaud, J.P.; Vaez, N. Optimization of silver nanoparticle biosynthesis by entomopathogenic fungi and assays of their antimicrobial and antifungal properties. J. Invertebr. Pathol. 2022, 190, 107749.
  16. Jamdagni, P.; Khatri, P.; Rana, J.S. Green synthesis of zinc oxide nanoparticles using flower extract of Nyctanthes arbor-tristis and their antifungal activity. J. King Saud Univ. Sci. 2018, 30, 168–175.
  17. Jamdagni, P.; Rana, J.S.; Khatri, P.; Nehra, K. Comparative account of antifungal activity of green and chemically synthesized zinc oxide nanoparticles in combination with agricultural fungicides. Int. J. Nano Dimens. 2018, 9, 198–208.
  18. Zhou, L.; Zhao, X.; Li, M.; Lu, Y.; Ai, C.; Jiang, C.; Liu, Y.; Pan, Z.; Shi, J. Antifungal activity of silver nanoparticles synthesized by iturin against Candida albicans in vitro and in vivo. Appl. Microbiol. Biotechnol. 2021, 105, 3759–3770.
  19. Shamraychuk, I.L.; Belyakova, G.A.; Eremina, I.M.; Kurakov, A.V.; Belozersky, M.A.; Dunaevsky, Y.E. Fungal proteolytic enzymes and their inhibitors as perspective biocides with antifungal action. Mosc. Univ. Biol. Sci. Bull. 2020, 75, 97–103.
  20. Padder, S.A.; Prasad, R.; Shah, A.H. Quorum sensing: A less known mode of communication among fungi. Microbiol. Res. 2018, 210, 51–58.
  21. Baier, F.; Tokuriki, N. Connectivity between catalytic landscapes of the metallo-β-lactamase superfamily. J. Mol. Biol. 2014, 426, 2442–2456.
  22. Lyagin, I.; Efremenko, E. Enzymes for detoxification of various mycotoxins: Origins and mechanisms of catalytic action. Molecules 2019, 24, 2362.
  23. Ayanwale, A.P.; Estrada-Capetillo, B.L.; Reyes-López, S.Y. Evaluation of antifungal activity by mixed oxide metallic nanocomposite against Candida spp. Processes 2021, 9, 773.
  24. Bhatt, V.K.; Patel, M.; Pataniya, P.M.; Iyer, B.D.; Sumesh, C.K.; Late, D.J. Enhanced antifungal activity of WS2/ZnO nanohybrid against Candida albicans. ACS Biomater. Sci. Eng. 2020, 6, 6069–6075.
  25. Abdelhamid, H.N.; Mahmoud, G.A.E. Antifungal and nanozyme activities of metal–organic framework-derived Appl. Organomet. Chem. 2023, 37, e7011.
  26. Pillai, A.M.; Sivasankarapillai, V.S.; Rahdar, A.; Joseph, J.; Sadeghfar, F.; Rajesh, K.; Kyzas, G.Z. Green synthesis and characterization of zinc oxide nanoparticles with antibacterial and antifungal activity. J. Mol. Struct. 2020, 1211, 128107.
  27. Arciniegas-Grijalba, P.A.; Patiño-Portela, M.C.; Mosquera-Sánchez, L.P.; Guerrero-Vargas, J.A.; Rodríguez-Páez, J.E. ZnO nanoparticles (ZnO-NPs) and their antifungal activity against coffee fungus Erythricium salmonicolor. Appl. Nanosci. 2017, 7, 225–241.
  28. Ilkhechi, N.N.; Mozammel, M.; Khosroushahi, A.Y. Antifungal effects of ZnO, TiO2 and ZnO-TiO2 nanostructures on Aspergillus flavus. Pestic. Biochem. Phys. 2021, 176, 104869.
  29. Miri, A.; Khatami, M.; Ebrahimy, O.; Sarani, M. Cytotoxic and antifungal studies of biosynthesized zinc oxide nanoparticles using extract of Prosopis farcta fruit. Green Chem. Lett. Rev. 2020, 13, 27–33.
  30. Parveen, S.; Wani, A.H.; Shah, M.A.; Devi, H.S.; Bhat, M.Y.; Koka, J.A. Preparation, characterization and antifungal activity of iron oxide nanoparticles. Microb. Pathog. 2018, 115, 287–292.
  31. Golipour, F.; Habibipour, R.; Moradihaghgou, L. Investigating effects of superparamagnetic iron oxide nanoparticles on Candida albicans biofilm formation. Med. Lab. J. 2019, 13, 44–50.
  32. Bouson, S.; Krittayavathananon, A.; Phattharasupakun, N.; Siwayaprahm, P.; Sawangphruk, M. Antifungal activity of water-stable copper-containing metal-organic frameworks. R. Soc. Open Sci. 2017, 4, 170654.
  33. Celis-Arias, V.; Loera-Serna, S.; Beltrán, H.I.; Álvarez-Zeferino, J.C.; Garrido, E.; Ruiz-Ramos, R. The fungicide effect of HKUST-1 on Aspergillus niger, Fusarium solani and Penicillium chrysogenum. New J. Chem. 2018, 42, 5570–5579.
  34. Veerana, M.; Kim, H.C.; Mitra, S.; Adhikari, B.C.; Park, G.; Huh, S.; Kim, S.; Kim, Y. Analysis of the effects of Cu-MOFs on fungal cell inactivation. RSC Adv. 2021, 11, 1057–1065.
  35. Tella, A.C.; Okoro, H.K.; Sokoya, S.O.; Adimula, V.O.; Olatunji, S.O.; Zvinowanda, C.; Ngila, J.C.; Shaibu, R.O.; Adeyemi, O.G. Synthesis, characterization and antifungal activity of Fe(III)metal–organic framework and its nano-composite. Chem. Afr. 2020, 3, 119–126.
  36. Wei, F.; Cui, X.; Wang, Z.; Dong, C.; Li, J.; Han, X. Recoverable peroxidase-like Fe3O4@MoS2-Ag nanozyme with enhanced antibacterial ability. Chem. Eng. J. 2021, 408, 127240.
  37. Liu, J.; Cheng, W.; Wang, Y.; Fan, X.; Shen, J.; Liu, H.; Wang, A.; Hui, A.; Nichols, F.; Chen, S. Cobalt-doped zinc oxide nanoparticle–MoS2 nanosheet composites as broad-spectrum bactericidal agents. ACS Appl. Nano Mater. 2021, 4, 4361–4370.
  38. Gow, N.A.R.; Latge, J.P.; Munro, C.A. The fungal cell wall: Structure, biosynthesis, and function. Microbiol. Spectr. 2017, 5, 10–128.
  39. Kühbacher, A.; Burger-Kentischer, A.; Rupp, S. Interaction of Candida species with the skin. Microorganisms 2017, 5, 32.
  40. Lyagin, I.; Stepanov, N.; Maslova, O.; Senko, O.; Aslanli, A.; Efremenko, E. Not a mistake but a feature: Promiscuous activity of enzymes meeting mycotoxins. Catalysts 2022, 12, 1095.
  41. Li, C.; Li, X.; Bai, C.; Zhang, Y.; Wang, Z. A chitinase with antifungal activity from naked oat (Avena chinensis) seeds. J. Food Biochem. 2019, 43, e12713.
  42. Dikbaş, N.; Uçar, S.; Tozlu, E.; Kotan, M.S.; ·Kotan, R. Antifungal activity of partially purified bacterial chitinase against Alternaria alternata. Erwerbs-Obstbau 2022.
  43. Zhang, W.; Ma, J.; Yan, Q.; Jiang, Z.; Yang, S. Biochemical characterization of a novel acidic chitinase with antifungal activity from Paenibacillus xylanexedens Z2–4. Int. J. Biol. Macromol. 2021, 182, 1528–1536.
  44. Rajninec, M.; Jopcik, M.; Danchenko, M.; Libantova, J. Biochemical and antifungal characteristics of recombinant class I chitinase from Drosera rotundifolia. Int. J. Biol. Macromol. 2020, 161, 854–863.
  45. Wang, N.-N.; Gao, K.-Y.; Han, N.; Tian, R.-Z.; Zhang, J.-L.; Yan, X.; Huang, L.-L. ChbB increases antifungal activity of Bacillus amyloliquefaciens against Valsa mali and shows synergistic action with bacterial chitinases. Biol. Control 2020, 142, 104150.
  46. Li, Q.; Hou, Z.; Zhou, D.; Jia, M.; Lu, S.; Yu, J. Antifungal activity and possible mechanism of Bacillus amyloliquefaciens FX2 against the postharvest apple ring rot pathogen. Phytopathology 2022, 112, 2486–2494.
  47. Lu, Y.; Wang, N.; He, J.; Li, Y.; Gao, X.; Huang, L.; Yan, X. Expression and characterization of a novel chitinase with antifungal activity from a rare actinomycete Saccharothrix yanglingensis Hhs.015. Protein Expr. Purif. 2018, 143, 45–51.
  48. Brzezinska, M.S.; Jankiewicz, U.; Kalwasinska, A.; Swiatczak, J.; Zero, K. Characterization of chitinase from Streptomyces luridiscabiei U05 and its antagonist potential against fungal plant pathogens. J. Phytopathol. 2019, 167, 404–412.
  49. Le, B.; Yang, S.H. Characterization of a chitinase from Salinivibrio sp. BAO-1801 as an antifungal activity and a biocatalyst for producing chitobiose. J. Basic Microbiol. 2018, 58, 848–856.
  50. Li, Z.; Xia, C.; Wang, Y.; Li, X.; Qiao, Y.; Li, C.; Zhou, J.; Zhang, L.; Ye, X.; Huang, Y.; et al. Identification of an endo-chitinase from Corallococcus sp. EGB and evaluation of its antifungal properties. Int. J. Biol. Macromol. 2019, 132, 1235–1243.
  51. Moon, C.; Seo, D.J.; Song, Y.S.; Hong, S.H.; Choi, S.H.; Jung, W.J. Antifungal activity and patterns of N-acetyl-chitooligosaccharide degradation via chitinase produced from Serratia marcescens PRNK-1. Microb. Pathog. 2017, 113, 218–224.
  52. Deng, J.-J.; Shi, D.; Mao, H.-H.; Li, Z.-W.; Liang, S.; Ke, Y.; Luo, X.-C. Heterologous expression and characterization of an antifungal chitinase (Chit46) from Trichoderma harzianum GIM 3.442 and its application in colloidal chitin conversion. Int. J. Biol. Macromol. 2019, 134, 113–121.
  53. Yilmaz, G.; Cadirci, B. Comparison of in vitro antifungal activity methods using Aeromonas sp. BHC02 chitinase, whose physicochemical properties were determined as antifungal agent candidate. Res. Sq. 2022.
  54. Sinitsyna, O.A.; Rubtsova, E.A.; Sinelnikov, I.G.; Osipov, D.O.; Rozhkova, A.M.; Matys, V.Y.; Bubnova, T.V.; Nemashkalov, V.A.; Sereda, A.S.; Tcsherbakova, L.A.; et al. Creation of chitinase producer and disruption of micromycete cell wall with the obtained enzyme preparation. Biochemistry 2020, 85, 717–724.
  55. Sachivkina, N.; Lenchenko, E.; Blumenkrants, D.; Ibragimova, A.; Bazarkina, O. Effects of farnesol and lyticase on the formation of Candida albicans biofilm. Vet. World 2020, 13, 1030–1036.
  56. Ling, L.; Cheng, W.; Jiang, K.; Jiao, Z.; Luo, H.; Yang, C.; Pang, M.; Lu, L. The antifungal activity of a serine protease and the enzyme production of characteristics of Bacillus licheniformis TG116. Arch. Microbiol. 2022, 204, 601.
  57. Deng, J.-J.; Huang, W.Q.; Li, Z.-W.; Lu, D.-L.; Zhang, Y.; Luo, X.-C. Biocontrol activity of recombinant aspartic protease from Trichoderma harzianum against pathogenic fungi. Enzyme Microb. Technol. 2018, 112, 35–42.
  58. Philip, N.V.; Koteshwara, A.; Kiran, G.A.; Raja, S.; Subrahmanyam, V.M.; Chandrashekar, H.R. Statistical optimization for coproduction of chitinase and beta 1,4-endoglucanase by chitinolytic Paenibacillus elgii PB1 having antifungal activity. Appl. Biochem. Biotechnol. 2020, 191, 135–150.
  59. Oyeleye, A.; Normi, Y.M. Chitinase: Diversity, limitations, and trends in engineering for suitable applications. Biosci. Rep. 2018, 38, BSR2018032300.
  60. Zhang, W.; Liu, Y.; Ma, J.; Yan, Q.; Jiang, Z.; Yang, S. Biochemical characterization of a bifunctional chitinase/lysozyme from Streptomyces sampsonii suitable for N-acetyl chitobiose production. Biotechnol. Lett. 2020, 42, 1489–1499.
  61. Li, S.; Zhang, B.; Zhu, H.; Zhu, T. Cloning and expression of the chitinase encoded by ChiKJ406136 from Streptomyces sampsonii (Millard & Burr) Waksman KJ40 and its antifungal effect. Forests 2018, 9, 699.
  62. Salazar, V.A.; Arranz-Trullén, J.; Navarro, S.; Blanco, J.A.; Sánchez, D.; Moussaoui, M.; Boix, E. Exploring the mechanisms of action of human secretory RNase 3 and RNase 7 against Candida albicans. Microbiol. Open 2016, 5, 830–845.
  63. Salazar, V.A.; Arranz-Trullén, J.; Prats-Ejarque, G.; Torrent, M.; Andreu, D.; Pulido, D.; Boix, E. Insight into the antifungal mechanism of action of human RNase N-terminus derived peptides. Int. J. Mol. Sci. 2019, 20, 4558.
  64. Tan, Y.; Ma, S.; Leonhard, M.; Moser, D.; Ludwig, R.; Schneider-Stickler, B. Co-immobilization of cellobiose dehydrogenase and deoxyribonuclease I on chitosan nanoparticles against fungal/bacterial polymicrobial biofilms targeting both biofilm matrix and microorganisms. Mater. Sci. Eng. C 2020, 108, 110499.
  65. Vidhate, R.P.; Bhide, A.J.; Gaikwad, S.M.; Giri, A.P. A potent chitin-hydrolyzing enzyme from Myrothecium verrucaria affects growth and development of Helicoverpa armigera and plant fungal pathogens. Int. J. Biol. Macromol. 2019, 141, 517–528.
  66. Silva, F.A.; Albuquerque, L.M.; Martins, T.F.; de Freitas, J.A.; Vasconcelos, I.M.; de Freitas, D.Q.; Moreno, F.B.M.B.; Monteiro-Moreira, A.C.O.; Oliveira, J.T.A. A peroxidase purified from cowpea roots possesses high thermal stability and displays antifungal activity against Colletotrichum gloeosporioides and Fusarium oxysporum. Biocatal. Agric. Biotechnol. 2022, 42, 102322.
  67. Zhang, L.; Tao, Y.; Zhao, S. A novel peroxiredoxin from the antagonistic endophytic bacterium Enterobacter sp. V1 contributes to cotton resistance against Verticillium dahliae. Plant Soil 2020, 454, 395–409.
  68. Pham, C.L.L.; de Francisco, B.R.; Valsecchi, I.; Dazzoni, R.; Pillé, A.; Lo, V.; Ball, S.R.; Cappai, R.; Wien, F.; Kwan, A.H.; et al. Probing structural changes during self-assembly of surface-active hydrophobin proteins that form functional amyloids in fungi. J. Mol. Biol. 2018, 430, 3784–3801.
  69. Valsecchi, I.; Dupres, V.; Stephen-Victor, E.; Guijarro, J.I.; Gibbons, J.; Beau, R.; Bayry, J.; Coppee, J.-Y.; Lafont, F.; Latgé, J.P.; et al. Role of hydrophobins in Aspergillus fumigatus. J. Fungi 2017, 4, 2.
  70. Valsecchi, I.; Lai, J.I.; Stephen-Victor, E.; Pillé, A.; Beaussart, A.; Lo, V.; Pham, C.L.L.; Aimanianda, V.; Kwan, A.H.; Duchateau, M.; et al. Assembly and disassembly of Aspergillus fumigatus conidial rodlets. Cell Surf. 2019, 5, 100023.
  71. Pham, C.L.L.; Rey, A.; Lo, V.; Soulès, M.; Ren, Q.; Meisl, G.; Knowles, T.P.J.; Kwan, A.H.; Sunde, M. Self-assembly of MPG1, a hydrophobin protein from the rice blast fungus that forms functional amyloid coatings, occurs by a surface-driven mechanism. Sci. Rep. 2016, 6, 25288.
  72. Saad, S.; Cereghetti, G.; Feng, Y.; Picotti, P.; Peter, M.; Dechant, R. Reversible protein aggregation is a protective mechanism to ensure cell cycle restart after stress. Nat. Cell Biol. 2017, 19, 1202e13.
  73. Beaussart, A.; Alsteens, D.; El-Kirat-Chatel, S.; Lipke, P.N.; Kucharíkova, S.; Dijck, P.V.; Dufrene, Y.F. Single-molecule imaging and functional analysis of Als adhesins and mannans during Candida albicans morphogenesis. ACS Nano 2012, 6, 10950–10964.
  74. Ho, V.; Herman-Bausier, P.; Shaw, C.; Conrad, K.A.; Garcia-Sherman, M.C.; Draghi, J.; Dufrene, Y.F.; Lipke, P.N.; Rauceo, J.M. An amyloid core sequence in the major Candida albicans adhesin Als1p mediates cell-cell adhesion. mBio 2019, 10, 10–128.
  75. Kumar, R.; Breindel, C.; Saraswat, D.; Cullen, P.J.; Edgerton, M. Candida albicans Sap6 amyloid regions function in cellular aggregation and zinc binding, and contribute to zinc acquisition. Sci. Rep. 2017, 7, 2908.
  76. Monniot, C.; Boisrame, A.; Costa, G.D.; Chauvel, M.; Sautour, M.; Bougnoux, M.-E.; Bellon-Fontaine, M.-N.; Dalle, F.; d’Enfert, C.; Richard, M.L. Rbt1 protein domains analysis in Candida albicans brings insights into hyphal surface modifications and Rbt1 potential role during adhesion and biofilm formation. PLoS ONE 2013, 8, e82395.
  77. Cabral, V.; Znaidi, S.; Walker, L.A.; Martin-Yken, H.; Dague, E.; Legrand, M.; Lee, K.; Chauvel, M.; Firon, A.; Rossignol, T.; et al. Targeted changes of the cell wall proteome influence Candida albicans ability to form single- and multi-strain biofilms. PLoS Pathog. 2014, 10, e1004542.
  78. Moreno-Ruiz, E.; Ortu, G.; de Groot, P.W.J.; Cottier, F.; Loussert, C.; Prevost, M.-C.; de Koster, C.; Klis, M.F.; Goyard, S.; d’Enfert, C. The GPI-modified proteins Pga59 and Pga62 of Candida albicans are required for cell wall integrity. Microbiology 2009, 155, 2004e20.
  79. Shanmugam, N.; Baker, M.O.; Ball, S.R.; Steain, M.; Pham, C.L.; Sunde, M. Microbial functional amyloids serve diverse purposes for structure, adhesion and defence. Biophys. Rev. 2019, 11, 287–302.
  80. Chernova, T.A.; Chernoff, Y.O.; Wilkinson, K.D. Yeast models for amyloids and prions: Environmental modulation and drug discovery. Molecules 2019, 24, 3388.
  81. Hirata, A.; Hori, Y.; Koga, Y.; Okada, J.; Sakudo, A.; Ikuta, K.; Kanaya, S.; Takano, K. Enzymatic activity of a subtilisin homolog, Tk-SP, from Thermococcus kodakarensis in detergents and its ability to degrade the abnormal prion protein. BMC Biotech. 2013, 13, 19.
  82. Dabbagh, F.; Negahdaripour, M.; Berenjian, A.; Behfar, A.; Mohammadi, F.; Zamani, M.; Irajie, C.; Ghasemi, Y. Nattokinase: Production and application. Appl. Microbiol. Biotechnol. 2014, 98, 9199–9206.
  83. Pilon, J.L.; Nash, P.B.; Arver, T.; Hoglund, D.; VerCauteren, K.C. Feasibility of infectious prion digestion using mild conditions and commercial subtilisin. J. Virol. Methods 2009, 161, 168–172.
  84. Saunders, S.E.; Bartz, J.C.; Vercauteren, K.C.; Bartelt-Hunt, S.L. Enzymatic digestion of chronic wasting disease prions bound to soil. Environ. Sci. Technol. 2010, 44, 4129–4135.
  85. McLeod, A.H.; Murdoch, H.; Dickinson, J.; Dennis, M.J.; Hall, G.A.; Buswell, C.M.; Carr, J.; Taylor, D.M.; Sutton, J.M.; Raven, N.D.H. Proteolytic inactivation of the bovine spongiform encephalopathy agent. Biochem. Biophys. Res. Commun. 2004, 317, 1165–1170.
  86. Dickinson, J.; Murdoch, H.; Dennis, M.J.; Hall, G.A.; Bott, R.; Crabb, W.D.; Penet, C.; Sutton, J.M.; Raven, N.D.H. Decontamination of prion protein (BSE301V) using a genetically engineered protease. J. Hosp. Infect. 2009, 72, 65–70.
  87. Yoshioka, M.; Miwa, T.; Horii, H.; Takata, M.; Yokoyama, T.; Nishizawa, K.; Watanabe, M.; Shinagawa, M.; Murayama, Y. Characterization of a proteolytic enzyme derived from a Bacillus strain that effectively degrades prion protein. J. Appl. Microbiol. 2007, 102, 509–515.
  88. Hui, Z.; Doi, H.; Kanouchi, H.; Matsuura, Y.; Mohri, S.; Nonomura, Y.; Oka, T. Alkaline serine protease produced by Streptomyces sp. degrades PrPSc. Biochem. Biophys. Res. Commun. 2004, 321, 45–50.
  89. Bahun, M.; Šnajder, M.; Turk, D.; Poklar Ulrih, N. Insights into the maturation of pernisine, a subtilisin-like protease from the hyperthermophilic archaeon Aeropyrum pernix. Appl. Environ. Microbiol. 2020, 86, e00971-20.
  90. Johnson, C.J.; Bennett, J.P.; Biro, S.M.; Duque-Velasquez, J.C.; Rodriguez, C.M.; Bessen, R.A.; Rocke, T.E. Degradation of the disease-associated prion protein by a serine protease from lichens. PLoS ONE 2011, 11, e19836.
  91. Chen, C.Y.; Rojanatavorn, K.; Clark, A.C.; Shih, J.C. Characterization and enzymatic degradation of Sup35NM, a yeast prion-like protein. Prot. Sci. 2005, 14, 2228–2235.
  92. Wang, J.J.; Borwornpinyo, R.; Odetallah, N.; Shih, J.C. Enzymatic degradation of a prion-like protein, Sup35NM-His6. Enzyme Microb. Technol. 2005, 36, 758–765.
  93. Sharma, R.; Gupta, R. Coupled action of γ-glutamyl transpeptidase-glutathione and keratinase effectively degrades feather keratin and surrogate prion protein, Sup 35NM. Biores. Tech. 2012, 120, 314–317.
  94. Rajput, R.; Gupta, R. Thermostable keratinase from Bacillus pumilus KS12: Production, chitin crosslinking and degradation of Sup35NM aggregates. Biores. Tech. 2013, 133, 118–126.
  95. Ningthoujam, D.S.; Mukherjee, S.; Devi, L.J.; Singh, E.S.; Tamreihao, K.; Khunjamayum, R.; Banerjee, S.; Mukhopadhyay, D. In vitro degradation of β-amyloid fibrils by microbial keratinase. Alzheimers Dement. 2019, 5, 154–163.
  96. Kim, N.; Lee, H.J. Redox-active metal ions and amyloid-degrading enzymes in Alzheimer’s disease. Int. J. Mol. Sci. 2021, 22, 7697.
  97. Manikandan, P.; Moopantakath, J.; Imchen, M.; Kumavath, R.; SenthilKumar, P.K. Identification of multi-potent protein subtilisin A from halophilic bacterium Bacillus firmus VE2. Microb. Pathog. 2021, 157, 105007.
  98. Kokwe, L.; Nnolim, N.E.; Ezeogu, L.I.; Sithole, B.; Nwodo, U.U. Thermoactive metallo-keratinase from Bacillus sp. NFH5: Characterization, structural elucidation, and potential application as detergent additive. Heliyon 2023, 9, e13635.
  99. Efremenko, E.; Aslanli, A.; Lyagin, I. Advanced situation with recombinant toxins: Diversity, production and application purposes. Int. J. Mol. Sci. 2023, 24, 4630.
  100. Efremenko, E.; Senko, O.; Stepanov, N.; Aslanli, A.; Maslova, O.; Lyagin, I. Quorum sensing as a trigger that improves characteristics of microbial biocatalysts. Microorganisms 2023, 11, 1395.
  101. Willaert, R.G. Adhesins of yeasts: Protein structure and interactions. J. Fungi 2018, 4, 119.
  102. Tian, X.; Ding, H.; Ke, W.; Wang, L. Quorum sensing in fungal species. Annu. Rev. Microbiol. 2021, 75, 449–469.
  103. Mehmood, A.; Liu, G.; Wang, X.; Meng, G.; Wang, C.; Liu, Y. Fungal quorum-sensing molecules and inhibitors with potential antifungal activity: A review. Molecules 2019, 24, 1950.
  104. Lee, K.; Lee, S.; Lee, S.H.; Kim, S.R.; Oh, H.S.; Park, P.K.; Choo, K.H.; Kim, Y.W.; Lee, J.K.; Lee, C.H. Fungal quorum quenching: A paradigm shift for energy savings in membrane bioreactor (MBR) for wastewater treatment. Environ. Sci. Technol. 2016, 50, 10914–10922.
  105. Ogawa, K.; Nakajima-Kambe, T.; Nakahara, T.; Kokufuta, E. Coimmobilization of gluconolactonase with glucose oxidase for improvement in kinetic property of enzymatically induced volume collapse in ionic gels. Biomacromolecules 2002, 3, 625–631.
  106. Aslanli, A.; Domnin, M.; Stepanov, N.; Efremenko, E. “Universal” antimicrobial combination of bacitracin and His6-OPH with lactonase activity, acting against various bacterial and yeast cells. Int. J. Mol. Sci. 2022, 23, 9400.
  107. Aslanli, A.; Domnin, M.; Stepanov, N.; Efremenko, E. Synergistic antimicrobial action of lactoferrin-derived peptides and quorum quenching enzymes. Int. J. Mol. Sci. 2023, 24, 3566.
  108. Hogan, D. Talking to themselves: Autoregulation and quorum sensing in fungi. Eukaryot. Cell 2006, 5, 613–619.
  109. Bu’LocK, J.D.; Jones, B.E.; Winskill, N. The apocarotenoid system of sex hormones and prohormones in mucorales. Pure Appl. Chem. 1976, 47, 191–202.
  110. Frolov, G.; Lyagin, I.; Senko, O.; Stepanov, N.; Pogorelsky, I.; Efremenko, E. Metal nanoparticles for improving bactericide functionality of usual fibers. Nanomaterials 2020, 10, 1724.
  111. Lyagin, I.; Stepanov, N.; Frolov, G.; Efremenko, E. Combined modification of fiber materials by enzymes and metal nanoparticles for chemical and biological protection. Int. J. Mol. Sci. 2022, 23, 1359.
  112. Lyagin, I.; Maslova, O.; Stepanov, N.; Presnov, D.; Efremenko, E. Assessment of composite with fibers as a support for antibacterial nanomaterials: A case study of bacterial cellulose, polylactide and usual textile. Fibers 2022, 10, 70.
  113. Lyagin, I.; Maslova, O.; Stepanov, N.; Efremenko, E. Degradation of mycotoxins in mixtures by combined proteinous nanobiocatalysts: In silico, in vitro and in vivo. Int. J. Biol. Macromol. 2022, 218, 866–877.
  114. Garces, F.; Fernández, F.J.; Montellà, C.; Penya-Soler, E.; Prohens, R.; Aguilar, J.; Baldomà, L.; Coll, M.; Badia, J.; Vega, M.C. Molecular architecture of the Mn2+-dependent lactonase UlaG reveals an RNase-like metallo-β-lactamase fold and a novel quaternary structure. J. Mol. Biol. 2010, 39, 715–729.
  115. González, J.M. Visualizing the superfamily of metallo-β-lactamases through sequence similarity network neighborhood connectivity analysis. Heliyon 2021, 7, e05867.
  116. Jang, E.-Y.; Son, Y.-J.; Park, S.-Y.; Yoo, J.-Y.; Cho, Y.-N.; Jeong, S.-Y.; Liu, S.; Son, H.-J. Improved biosynthesis of silver nanoparticles using keratinase from Stenotrophomonas maltophilia R13: Reaction optimization, structural characterization, and biomedical activity. Bioprocess Biosyst. Eng. 2018, 41, 381–393.
  117. Abo-Zaid, G.; Abdelkhalek, A.; Matar, S.; Darwish, M.; Abdel-Gayed, M. Application of bio-friendly formulations of chitinase-producing Streptomyces cellulosae Actino 48 for controlling peanut soil-borne diseases caused by Sclerotium rolfsii. J. Fungi 2021, 7, e167.
  118. Gaonkar, S.K.; Furtado, I.J. Biorefinery-fermentation of agro-wastes by Haloferax lucentensis GUBF-2 MG076878 to haloextremozymes for use as biofertilizer and biosynthesizer of AgNPs. Waste Biomass Valorization 2022, 13, 1117–1133.
  119. Guilger-Casagrande, M.; Germano-Costa, T.; Pasquoto-Stigliani, T.; Fraceto, L.F.; de Lima, R. Biosynthesis of silver nanoparticles employing Trichoderma harzianum with enzymatic stimulation for the control of Sclerotinia sclerotiorum. Sci. Rep. 2019, 9, e14351.
  120. Guilger-Casagrande, M.; Germano-Costa, T.; Bilesky-José, N.; Pasquoto-Stigliani, T.; Carvalho, L.; Fraceto, L.F.; de Lima, R. Influence of the capping of biogenic silver nanoparticles on their toxicity and mechanism of action towards Sclerotinia sclerotiorum. J. Nanobiotechnol. 2021, 19, e53.
  121. Dror, Y.; Ophir, C.; Freeman, A. Silver-enzyme hybrids as wide-spectrum antimicrobial agents. In Innovations and Merging Technologies in Wound Care; Gefen, A., Ed.; Elsevier Inc.: Amsterdam, The Netherlands, 2020; pp. 293–307.
  122. Xia, S.; Xu, Y.; Hoy, R.; Zhang, J.; Qin, L.; Li, X. The notorious soilborne pathogenic fungus Sclerotinia sclerotiorum: An update on genes studied with mutant analysis. Pathogens 2020, 9, e27.
  123. Chet, I.; Henis, Y. Effect of catechol and disodium EDTA on melanin content of hyphal and sclerotial walls of Sclerotium rolfsii sacc. and the role of melanin in the susceptibility of these walls to β-(1→3) glucanase and chitinase. Soil Biol. Biochem. 1969, 1, 131–138.
  124. Melo, B.S.; Voltan, A.R.; Arruda, W.; Lopes, F.A.C.; Georg, R.C.; Ulhoa, C.J. Morphological and molecular aspects of sclerotial development in the phytopathogenic fungus Sclerotinia sclerotiorum. Microbiol. Res. 2019, 229, e126326.
  125. Yu, D.; Wang, Y.; Zhang, J.; Yu, Q.; Liu, S.; Li, M. Synthesis of the ternary nanocomposites composed of zinc 2-methylimidazolate frameworks, lactoferrin and melittin for antifungal therapy. J. Mater. Sci. 2022, 57, 16809–16819.
  126. Babina, S.E.; Kanyshkova, T.G.; Buneva, V.N.; Nevinsky, G.A. Lactoferrin is the major deoxyribonuclease of human milk. Biochemistry 2004, 69, 1006–1015.
  127. Fernandes, K.E.; Weeks, K.; Carter, D.A. Lactoferrin is broadly active against yeasts and highly synergistic with amphotericin B. Antimicrob. Agents Chemother. 2020, 64, e02284-19.
Subjects: Microbiology
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to : , , , , , ,
View Times: 402
Revisions: 2 times (View History)
Update Date: 24 Jul 2023