Fungal Surfactants and Emulsifiers Enhance Pollutant Partition into Aqueous Phases

Hydrophobins and similar fungal proteins can affect the bioavailability of hydrophobic contaminants in the soil by promoting their dispersion in the aqueous phase. Indeed, a partial solubilization of hydrophobic drugs and increase of their bioavailability have been achieved using the SC3 hydrophobin from the Basidiomycete SchizophyUum commune (Haas Jimoh Akanbi et al. 2010), and possible hydrophobin-like proteins able to solubilize PAH have been found in Aspergillus brasiliensis (Sanchez-Vazquez et al. 2018). Due to these versatile surface-active properties, hydrophobins have been proposed as a tool for oil recovery (Blesic et al. 2018).

In addition to hydrophobins, fungi produce a wide range of biosurfactants and emulsifiers with diverse chemical natures (Table 2). Although both categories overlap to some extent, surfactants are molecules able to lower interfacial tensions between water and air or organic phases, while emulsifiers are defined by the ability to stabilize oil-in-water emulsions. Most of these molecules or molecular complexes can be classified in three broad categories: glycoproteins/glycopeptides, lipoproteins/lipopeptides, and glycolipids. Biosurfactants from fungi, and especially yeast and yeast-like strains, have been known for decades, yet until recently little information was available about biosurfactants from filamentous fungi (Bliardwaj 2013). However, as the demand for bio-sourced and biodegradable compounds in general increased over the last 10 years, microbial surfactants became a growing field of research.

Based on the overview presented in Table 2, glycolipids appear to be the most widespread among fungal surfactants and emulsifiers. The high molecular weight glycoprotein liposan from Candida lipolytica (Cirigliano and Carman 1985), sophorolipids from Starmerella (Candida) bombicola (Cooper and Paddock 1984), and ustilagic acid, a cellobiose lipid from the Zygomycete Ustilago maydis (Frautz et al. 1986), were the first main biosurfactants characterized in fungi. Mannosylerythritol lipids (MEL) produced by several smut fungi (Arutchelvi et al. 2008) and polyol lipids found in Aureobasidium pullulans and oleaginous yeasts of the Rhodotorula genus (Garay et al. 2018) are two other more recently identified classes of fungal surfactants. Several functions have been proposed for these extracellular surfactants: in a similar way to hydrophobins, they are likely involved in regulating the interaction of hyphae with surfaces. Polyol lipids and sophorolipids in particular may favor fungal growth on plant tissues by assisting the breakdown of cuticular waxes. In addition, other functions have been identified: the mobilization of hydrophobic substrates, and defense against competing microorganisms through antibacterial or antifungal properties (Puchkov et al. 2002, Arutchelvi et al. 2008, Garay et al. 2018). Some of these molecules appear to be produced constitutively, while others are induced by cultivation in presence of oil, and may be derivatives of lipid catabolism.

The potential industrial applications of biosurfactants was the main driver for research rather than the understanding of fungal physiology and ecology, hence the limited knowledge of their biological function in nature. Interestingly, many surfactant-producing fungal strains were identified as such while investigating their potential for the remediation of hydrocarbons and other hydrophobic soil contaminants (Batista et al. 2010, de Luna et al. 2015, Azin et al. 2018, Zadeh et al. 2018, do Amaral Marques et al. 2019, Pele et al. 2019), including PAH (Deziel et al. 1996, Nikiforova et al. 2009, Deng et al. 2010, Veignie et al. 2012). Rafin et al. showed an increase in Benzo[a]pyrene (BaP) concentration over time in fungal culture filtrates, suggesting the production of extracellular mobilizing agents able to partially stabilize BaP in the water phase (Rafin et al. 2013, Fayeulle et al. 2019). There is also evidence of emulsification of aliphatic and aromatic hydrocarbons by Penicillium citrinum (Camargo-de-Morais et al. 2003). Complexation agents are also produced by some strains known for biodegradation of hydrocarbons such as Fusarium solani (Veignie et al. 2012).

Remediation enhancement in presence of surfactant is likely driven by two main mechanisms involving i) solubilization and transport of organic compounds into micelles, and ii) displacement of entrapped NAPLs due to interfacial tension reduction (Paria 2008). Indeed, when in presence of surfactant, the fraction of HOC found in micellar form is directly available for degrading bacteria, in a similar way to the dissolved fraction of HOC (Brown 2007). Biosurfactant- enhanced desorption of phenanthrene from suspended clays and humic acid was observed by Garcia-Junco (Garcia-Junco et al. 2003). Similar mechanisms may be involved for fungal surfactants, increasing the bioavailability of HOC in soils to both the fungus itself and other microorganisms.

Alteration of the Soil Matrix and Effect on the Retention of Hydrophobic Pollutants

Soil structure is determined by the aggregation of its components, including minerals (clays and other silicates, calcium carbonate, metallic oxides) and organic matter. The relative content in particles of various size ranges defines soil texture, regardless of their chemical nature. Fungi greatly contribute to shaping the soil matrix by simultaneously promoting aggregate formation, and altering minerals and organic matter (Gadd et al. 2012, Ritz and Young 2004).

Table 2 Fungal strains identified as sources of extracellular biosurfactants or bioemulsifiers, and chemical nature thereof (when characterized)




Cunninghamella echinulata


Andrade Silva et al. 2014

Mucor circinelloides

lipopeptid glycolipid

do Amaral Marques

et al. 2019, Zadeli et al. 2018

Mucor indicus


Oje et al. 2016

Rhizopus airhizus


Pele et al. 2019

Cunninghamella echinulata


Andrade Silva et al. 2014

Candida glabrata


de Luna et al. 2009

Candida tropicalis


Batista et al. 2010

Dipodascus (Candida) ingens

(fatty acids)

Amezcua-Vega et al. 2007

Kluyveromyces lactis (Candida sphaerica)


de Luna et al. 2015

Kluyveromyces marxianus


Lukondeh et al. 2003

Lachancea thermotolerans


Mousavi et al. 2015

Nakazawaea (Candida) ishiwadae


Thanomsub et al. 2004

Saccharomyces cerevisiae

parietal mannoprotein

Cameron et al. 1988

Starmerella (Candida) bombicola


Cooper and Paddock 1984

Wickerhamomyces anomalus


Texeira Souza et al. 2017

Wickerhamiella domercqiae


Ma et al. 2014

Yarrowia (Candida) lipolytica

high molecular

weight glycoprotein “Liposan”

Cirigliano and Carman 1985

Fusarium fujikuroi


Loureiro dos Reis et al. 2018

Fusarium neocosmosporiellum unknown

Azin et al. 2018

Fusarium proliferation

fatty amide

Bhardwaj et al. 2015

Fusarium solani


Yeignie et al. 2012

Fusarium sp.


Sena et al. 2018

Phialemonium sp.


Guimaraes Martins et al. 2006

Penicillium chrysogenum


Gautam et al. 2014

Penicillium citrinum


Camargo-de-Morais et al. 2003

Penicillium sp.

Lipids, carbohydrates, protein

Luna-Л elasco et al. 2007

Penicillium sp.


Sena et al. 2018

Aspergillus flavus

phenyl glycoside

Ishaq et al. 2015

Aspergillus fumigatus


Guimaraes Martins et al. 2006

Aspergillus ustus


Kiran et al. 2009

Aspergillus niger


Costa Sperb et al. 2018

Aureobasidium pullulans

polyol lipids “liamocins”

Price et al. 2013

Aureobasidium thailandense

fatty acid ester

Meneses et al. 2017

Exophiala dermatitidis


Chiewpattanakul et al. 2010

Trichoderma sp.


Sena et al. 2018

Table 2 Contd...

Table 2 (contd...) Fungal strains identified as sources of extracellular biosurfactants or bioemulsifiers, and chemical nature thereof (when characterized)





“white rots”

Ceriporia lacerata

mannosylerythritol lipids

Niu et al. 2017

Pleurotus djamor


Yelioglu and Urek 2016

PJeurotus ostreatus


Nikiforova et al. 2009

Trametes versicolor


Lourenco et al. 2018


Pseudozyma spp.

mannosylerythritol lipids cellobiose lipids

Kitamoto et al. 1993, Morita et al. 2007, Morita et al. 2013

Sporisorium sp.

mannosylerythritol lipids

Alimadadi et al. 2018

Ustilago maydis

cellobiose lipid «ustilagic acid»

Frautz et al. 1986

Ustilago spp.

mannosylerythritol lipids

Spoeckner et al. 1999, Morita et al. 2008, 2009

Oleaginous yeasts

Cutaneotrichosporon curvatun (Cryptococcus curvatus)


Ma et al. 2014

Rhodotorula babjevae


Sen et al. 2017

Rhodotorula spp.

polyol lipids

Garay et al. 2018

Vanrija (Cryptococcus) humicola

cellobiose lipid “mycocin”

Puchkov et al. 2002

In combination with organic matter content, porosity is one of the main parameters affecting the retention of contaminants. Indeed, small pores and high total pore volume mean a high surface available for adsorption, and inaccessibility to degrading organisms and solutions (Ren et al. 2018, Yu et al. 2018). HOC are known to have a greater affinity for fine particles, and tend to sorb onto clay minerals, which exhibit hydrophobic surfaces in a layered structure (Jaynes and Boyd 1991, Yu et al. 2018). As a result, pollutant retention is usually higher in soils with a finer texture (Amellal et al. 2001).

Filamentous fungi develop extensive hyphal networks in the soil and can penetrate solid materials while exploring the substrate. Growing hyphal tips are subjected to swelling/shrinking cycles due to changes in intracellular osmotic pressure, exerting a mechanical force on the surfaces encountered, and cause biomechanical weathering of minerals through penetration into cracks and micropores (Fomina et al. 2006). Fungal weathering is also biochemical: fungi acidify their immediate surroundings by excreting organic acids (acetic, citric, oxalic, formic acid) present in hyphal exsudates, as well as dissolved respiratory C02, thus promoting mineral dissolution. Zhang et al. (2016) suggest that organic acids produced through the metabolization of alkanes could in turn enhance the porosity of the solid matrix by dissolving carbonated minerals, thus increasing the accessibility of oil deposits. Metal chelators including siderophores and phenols are also involved in the chemical alteration of soil minerals, destabilizing their chemical structure by solubilizing the metallic elements (Gadd et al. 2012). Such chemical and physical alteration of minerals can enhance the release of inaccessible contaminant droplets or particles trapped in the soil solid matrix. Erosion phenomena lead to the fragmentation of the solid substrate into smaller particles or colloids susceptible to be carried through soil pores by water flows. Colloid-facilitated transport may contribute to the dispersion of sorbed contaminants in soils (de Jonge et al. 2004). Indeed, some studies show that transport in a particle-bound form account for a significant proportion of the leaching of hydrophobic pesticides through macropores (Villholth et al. 2000, Kjaer et al. 2011).

One of the major functions of saprotrophic fungi affecting soil structure and chemical cycles is the biodegradation of dead organic matter. Fungi secrete a wide range of extracellular lytic enzymes including cellulases, xylanases, lipases, proteases, and lignin-degrading enzymes in the case of white rots. Through the enzymatic hydrolysis and dissolution of complex macromolecules, fungi disrupt organic matter aggregates, rendering bound HOC more accessible due to two complementary effects: the removal of physical hindrances preventing microorganisms from accessing the source of pollutant, and the release of mobile organic matter fragments promoting the dispersal of sorbed HOC. Soluble humic acids act as carriers increasing the mobility and bioacessibility of HOC despite very low aqueous concentrations and making them directly available for uptake even without desorption and solubilization in the water phase (Smith et al. 2009). Hydrophobic contaminants tend to sorb onto suspended humic acid-clay complexes (Garcia-Junco et al. 2003). Moreover, the biodegradation of organic matter by fungi also increases the dissolved organic carbon content, which has been linked to facilitated diffusion of HOC from NAPLs to the water phase (Smith et al. 2011).

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