ORIGINAL ARTICLE

The role of ethanol in preventing biofilm formationof Penicillium purpurogenum

Sherif M. Husseiny & Hussein Abd El Kareem &

Ola M. Gomaa & Riham Talaat

Received: 25 July 2013 /Accepted: 8 December 2013# Springer-Verlag Berlin Heidelberg and the University of Milan 2013

Abstract The use of fungi in biotechnology requires that nocell loss takes place; a maximal level of cell–nutrient interac-tion is required to achieve efficient performance. The occur-rence of high cell densities or loss of biomass through cell–surface interaction prevents the desired result. The main pur-pose of adding ethanol was to manipulate the cell–cell andcell–surface adhesion through manipulating cell surface prop-erties. Scanning electron microscopy indicated that the type ofsurface and its treatment with ethanol controls the adhesionand biofilm formation of Penicillium purpurogenum. Gammairradiation slightly affected the wettability of polystyrenestrips at 0.5 and 1 kGy, thus slightly decreasing the adhesion,but was not as effective as using ethanol to control the adhe-sion. The presence of ethanol in the media caused a decreasein surface-bound proteins from 0.348 to 0.133 mg/ml, whilesurface exopolysaccharides showed a minimal decrease.Ethanol induced oxidative stress which reached its peak when2.5 % v/v ethanol was added to the media; this was represent-ed by both intracellular and extracellular catalase and lipidperoxidation. On the other hand, fungal biomass and pigmentshowed a decrease as the ethanol concentrations increased.Therefore, ethanol could be employed to control the surfaceproperties of a fungus, and to inhibit biofilm formation toobtain a high surface area for the fungus to be employed inany biotechnological process.

Keywords Penicillium purpurogenum . Adhesion . Biofilmformation . Ethanol . Oxidative stress response

Introduction

The adherence of microbial cells onto surfaces often results ina build-up of aggregates and the formation of what is knownas “biofilm”. Biofilm formation is the oldest and most pow-erful form of life; its strength arises from the microbial cells’ability to produce layers of extracellular polymeric substancewhich offer protection against biocides and toxins (Stoodleyet al. 2004). However, as crucial as it is for microorganisms toform biofilms to protect their integrity and continue theirsurvival in any given harsh environment, biofilm formationis considered a common threat in fields such as the foodindustry (Simões et al. 2010) and the biomedical field (Haoet al. 2012). It is also perceived as a threat in wastewatertreatment reactors for their ability to cause corrosion, odor,and hydrogen sulfide (Jiang and Yuan 2013). Biofilms mayact as a harbor for pathogenic microbial cells in drinking waterreservoirs (Piriou et al 1997). Problems arising from biofilmformation is due to the cost associated with the losses itcauses: the deterioration in plant performance, the decreasein the quality and quantity of the product, the damage of theconstructing material, and the cost of cleaning processes orcost of addition al biocides or labor used to replace or clean thetanks (Al-Juboori and Yusaf 2012). There are some com-pounds that, when added to the medium, prevent cell adhe-sion, hence preventing cell aggregation and biofilm formationsuch as biosurfactants (Monteiro et al 2011), dipeptide cis-cyclo(Leucyl-Tyrosyl) (Scopel et al 2013), or antibiotics(Ferrnandez-Olmos et al. 2012). Other methods of controllingbiofilm formation include membrane surface modification orbiochemical techniques which involve degrading the EPSusing enzymes, bacteriophages, and signaling proteins(Al-Juboori and Yusaf 2012). The attachment process betweenfungal spores and/or hyphae and substrates is considered avery complex process; it mainly depends on the physicochem-ical surface interaction; specific molecular factors being

S. M. HusseinyBotany Department, Girl’s College, Ain Shams University, Cairo,Egypt

H. A. El Kareem :O. M. Gomaa (*) :R. TalaatMicrobiology Department, National Center for Radiation Researchand Technology (NCRRT), Cairo, Egypte-mail: ola_gomaa@hotmail.com

Ann MicrobiolDOI 10.1007/s13213-013-0788-5

glycoproteins, hydrophobins, carbohydrates, and lipids(Priegnitz et al 2012).

Biofilm removal or prevention is considered somewhateasier when there is no need for live microbial cells, but it isvery difficult to prevent its formation and still keep the cellslive and intact. Ethanol is an agro-industrial waste that isproduced in huge quantities in Egypt in the process of makingsugar from sugar cane; therefore, it is very cheap and abun-dant. It is an aliphatic alcohol that has been used in variousways, when added at 70 % (w/v) it is employed as a disinfec-tant, while at 5–10 % it is bacteriostatic (Sissons et al 1996). Itcan be used as a carbon source for fungi (Mogensen et al2006), as a supplemental electron donor to stimulate microbialreduction of nickel and iron (Akob et al 2008), and as aninducer for laccase (phenol oxidase) enzyme in white rot fungi(Alves et al 2004). The fungus Penicillium purpurogenum is afilamentous fungi which belongs to the phylum Ascomycota,and is known for its biotechnological applications in industry:it is known to produce red pigment (Mendez et al 2011), and ithas also been characterized as phenol oxidase producer in ourlaboratory (data unpublished). In the following work, ethanolwill be employed to control the adhesion and biofilm forma-tion of Penicillium purpurogenum, through examining someof the parameters controlling microbial adhesion.

Materials and methods

Fungal isolation and cultivation conditions

The fungus used was isolated from soil about 40 km outsideCairo, Egypt. About 10 g were added to 90 ml sterile salinesolution and shaken for 1 h. After serial dilution, 0.1 ml of theappropriate dilution was spread over Czapek’s Yeast agar(CZYA) plates; the media consisted of the following per L:K2HPO4 1 g, yeast extract 5 g, sucrose 30 g, Czapek’s con-centrate 10 ml, and agar 20 g. The Czapek’s concentrate wascomposed of the following per L: MgSO4·7H2O 5 g, NaNO3

30 g, KCl 5 g, FeSO4·7H2O 0.1 g, ZnSO4·7H2O 0.1 g, andCuSO4·5H2O 0.05 g. After 7 days incubation, the sampleswere purified by streaking onto clean CZYA plates. Periodicalsubculturing of fungi was performed on agar slants and storedat 4 °C. The preliminary identification of the isolates was doneon water agar plates based on their morphology according toPitt and Hocking (1985).

Morphological study

Using CZYA plates with and without ethanol, a glass coverslide was placed at an angle of 45°, the fungus was inoculatedat the base of the cover slide, andwas left to incubate for 7 days.The cover slides with the grown fungus on the edge were usedfor scanning electron microscopy as described below.

Adhesion and biofilm formation

Polystyrene sheet, tin sheet, and glass cover slides were all cutinto 0.25×0.5 cm strips. About 100 μl of spore suspensionwas added to the wells of a round-bottom 96-well microtiterplate, and the wells were divided to groups, each containingstrips of polystyrene, tin, or glass, with and without ethanol.Another group was used with ethanol-immersed strips. Theplates were left to incubate for 24 h as previously described.The strips were taken from the wells with a sterile forceps andleft to dry in air. Scanning electron micrographs of the adhe-sion on the strips was carried out using a JOEL JMS 5600scanning electron microscope; after the strips were air-dried,they were glued separately on to brass stubs using double-sided adhesive tape and were coated with a thin layer of goldunder reduced pressure. The images were captured at magni-fications of ×750 using an electron beam high voltage of30 kV.

Gamma radiation

Polystyrene strips were placed each in separate pouches andwere used for gamma radiation experiments. Gamma irradia-tion was performed in triplicates at the cobalt source located atNCRRT, Cairo, Egypt. The strips were subjected to the fol-lowing doses: 0.3, 0.5, and 1 kGy at a dose rate of 2.95 kGy/h.The doses employed were chosen based on a series of exper-iments to ensure that the shape and properties of the polymerdid not change (data unpublished). Fungal spore suspension,incubation in microtitre plates, and scanning electron micros-copy were performed as previously mentioned.

Biochemical assays

A single 4 mm plug cut from the periphery of a 7-day-oldculture was taken used the broad side of a sterile tip was usedto inoculate a set of cultures. Ethanol was added under sterile -conditions to different CZY liquid cultures in 100-mlErlenmeyer flasks with 20 ml working volume on the day ofinoculation to obtain final concentrations of 0, 2.3, 5, 7.5, and10 % v/v. The Erlenmeyer flasks were incubated under staticconditions at 30 °C for 7 days. The cultures were used for thefollowing biochemical assays.

Catalase

Catalase was measured according to the method of Beers andSizer (1952). The disappearance of peroxide was followedspectrophotometrically at 240 nm using a Schimadzu UV2100 spectrophotometer. One Unit was defined as the quantityof catalase that decomposes 1 µmol of H2O2 per min at 25 °C(pH 7.0). The reaction mixture consisted of 0.05 M potassium

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phosphate buffer (pH 7) containing 0.059 M hydrogenperoxide.

Lipid peroxidation

Lipid peroxidation was calculated as the concentration ofmalondialehyde (MDA) (the end product of lipid peroxida-tion) in the cell wall of pellets of copper-free and copper-amended cultures. Lipid peroxidation was determined as thio-barbituric acid reactive substance (TBARS) according toYoshika et al. (1979).

Mycelial weight

The fungal biomass obtained at the end of incubation periodwas washed with distilled water and dried in an oven at 70 °Cfor 24 h. Dry biomass was determined as dry weight pervolume.

Red pigment assay

Red pigment assay was performed using the extracellu-lar fluid (ECF) for each culture; the ECF were used forvisible spectrophotometric analysis at 492 nm to test thechanges in color for all tested ethanol concentrations(Mendez et al 2011).

Exopolysaccharides (EPS)

Cultures in the previous experiment were centrifuged at5,000 rpm for 15 min, the supernatant was removed and95 % ethanol was added to the cells and incubated at 4 °Covernight to release surface-bound exopolysaccharides(Nehad and El-Shamy 2010), while soluble EPS was deter-mined in the culture supernatant directly. Both surface-bound

and soluble EPS were determined using the phenol-sulfuricmethod (Chaplin and Kennedy 1986), absorbance was mea-sured at 490 nm, glucose was used as standard.

Surface-bound protein

The surface-bound proteins were extracted according to amodified method of Castellanos et al. (1997), the cells wereharvested, washed twice with PBS, and the pellets resuspend-ed in 10 ml 6 M urea for 90 min at 22 °C. The cell suspensionwas centrifuged at 1,600 g for 10 min at 10 °C, and thesupernatant was used to detect the protein content usingLowry’s method (Lowry et al. 1951) using bovine serumalbumin (BSA) as a standard.

Cell surface charge

Spore suspension of Penicillium purpurogenum was used todetect the cell surface charge in the presence (2.5 and 5 % v/v)and absence of ethanol using the two-phase partitioning assayas described by Castellanos et al (1997). Each system wasdone separately; the phases 1 and 2 were added consecutively.System I consisted of 7.13 % polyethylene glycol (PEG) in150 mM NaCl as phase 1 and 8.75 % dextran in 150 mMNaCl as phase 2, while system II consisted of 7.13 % PEG in150 mM as phase 1 and 8.35 % dextran and 0.4 % dextransulfate in 150 mM NaCl as phase 2. The results wereexpressed as Δ log G which is defined by the followingequation:

Δ log G ¼ log G value for system II=G value for system Ið Þ

Where G = % cells in top phase/ % cells in the rest of thesystem. Values larger than zero indicate negatively chargedcell surface.

Control Penicillium purpurogenum culture Ethanol amended Penicilliumpurpurogenum culture

Fig. 1 Scanning electronmicrographs of ethanol amendedPenicillium purpurogenumcultures as compared to controlcultures, magnification ×2000

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Results

Morhological changes

Cultures of Penicillium purpurogenum were examinedfor morphological changes after growth in ethanol andin control cultures. The pictures in Fig. 1 clearly showthat ethanol-amended cultures exhibited la oose myce-lial network as compared to a tight network in controlcultures.

To study the effect of ethanol on biofilm formation ondifferent substrates, glass, polystyrene, and tin foil strips wereused, each categorized to control and ethanol-amendedgroups. Figure 2 shows that the different cultures grown inethanol showed obviously less adhesion on different media ascompared to those grown in ethanol-free media. The leastgrowth was exhibited on glass strips, followed by polystyrenestrips, and tin foil strips.

The manipulation of the cell surface plays a role in adhe-sion, as when gamma radiation was used in different doses on

Fig. 2 Penicilliumpurpurogenum grown on glass (1,2), polystyrene strips (3, 4) and tinfoil strips (5, 6) in control andethanol amended microtitreplates, respectively

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polystyrene strips and Penicillium purpurogenum was incu-bated with gamma irradiated strips, the results show an in-crease in the loose mycelial network which increased at 0.3and 0.5 kGy and was maximal at 1 kGy (Fig. 3).

Biological changes

The results in Fig. 4 clearly show that the addition of ethanolto the culture media results in a parallel increase in bothextracellular and intracellular catalase; the former is producedin quantity. Catalase activity reached its peak at 6.49 U/mlwhen 2.5 % v/v was added to the media, after which theactivity showed a gradual decrease and reached its minimumof 2.49 U/ml when 10 % v/v ethanol was added to the media.Although intracellular catalase followed the same pattern, thevalues were below those shown for extracellular catalase.

Another indication of stress by ethanol is shown in Fig. 5which represents the degree of lipid peroxidation and mycelialgrowth in the presence of different concentrations of ethanol.The figure shows that lipid peroxidation reached its maximumof 0.23 mg/mg mycelia when 2.5 % v/v ethanol was added tothe culture medium, above which there was a drop in lipidperoxidation. On the other hand, mycelial growth was main-tained in the presence of 2.5 and 5 % v/v ethanol along withcontrol cultures and was represented as 100 % mycelial

growth, above which there was a sharp decrease in mycelialgrowth that reached only 5 %.

Penicillium purpurogenum produces a red pigment, whichwas affected by the addition of ethanol to the culture medium.The deep rich red color of control cultures showed absorbanceof 5.8 but exhibited a lighter shade as the ethanol concentra-tion increased, until it reached an absorbance of 0.97 when10 % v/v ethanol was added to the culture medium (Fig. 6).

Penicillium purpurogenum adhesion on non-irradiated polystyrene strip

Penicillium purpurogenum adhesion on 0.3 kGy irradiated polystyrene strip

Penicillium purpurogenum adhesion on 0.5 kGy irradiated polystyrene strip

Penicillium purpurogenum adhesion on 1 kGy irradiated polystyrene strip

Fig. 3 Effect of gamma radiationon adhesion of Penicilliumpurpurogenum on polystyrenestrips grown in microtiter plates

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Fig. 4 Intracellular and extracellular catalase in Penicilliumpurpurogenum grown in cultures amended with different ethanolconcentrations

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One of the important factors which control biofilm forma-tion is the surface exopolysaccharides and surface-bound pro-teins. Figure 7 shows that there is no distinct change in EPS atdifferent ethanol concentrations, the results varying from 10.3to 9.5 mg/ml. On the other hand, the surface-bound proteinsexhibited a drop which reached 0.133 mg/ml as compared to0.348 mg/ml in control cultures which means there was about2.6-fold drop in surface-bound proteins when ethanol wasadded to the culture medium.

To study the degree of cell attachment in the presence ofethanol, the cell surface charge (CSC) was calculated in thepresence and absence of ethanol. Figure 8 shows an increasein CSC value from 0.043 in control cultures, to 0.18 for2.5 % v/v ethanol, and reached a maximum value of 0.32when 5 % v/v ethanol was added to the culture media.

Discussion

Biofilm formation is considered a complex process; it isinitiated by the adhesion of cells to cells or cells to surface.

The adhesion process is comprised of two parts: an initialphysical attachment takes place first; this is due to weak,reversible van der Waals forces. Later on, an irreversiblestrong adhesion follows as a result of either a ligand-receptor or extracellular polysaccharides (Amiri et al 2005).To test the fungal adhesion and biofilm formation, scanningelectron microscopy was used as a monitoring technique(Al-Juboori and Yusaf 2012). The results show thatfungal morphology in the presence of ethanol was markedlyaffected; control cultures showed tightly bound mycelialgrowthwhile ethanol-amended cultures showed a loose boundmycelial growth. This result is another confirmation that thepresence of non-growth-inhibiting ethanol in the medium hadan effect on the surface characteristics and architecture of thegrowing mycelium. For detection of fungal adhesion on dif-ferent substrates, scanning electron microscopy was also usedto monitor the adhesion on glass, polystyrene, and tin foil inthe presence and absence of ethanol. The results show thatfungi showed less adhesion in the presence of ethanol for allthree substrates used; however, there was also a variation inthe adhesion even when ethanol was added, the least being onglass, followed by polystyrene, and the highest adhesion beingon tin foil. Glass is stated to be a wettable surface whilepolystyrene is a less wettable surface (Amiri et al 2005).This suggests that the adhesion is related to both the changeson the cell surface and the hydrophobicity of the substrate.This line of evidence is further supported by the fact that usinggamma radiation to vary the wettability of polystyrene sur-faces caused an alteration in the adhesion; the same tookplace when polystyrene sheets were immersed in etha-nol. The use of radiation to decrease the adhesion of awettable substrate has been used before: UV radiation wasused to decrease polystyrene wettability and hence decreasethe adhesion of Penicillium expansum (Amiri et al 2005).

The most two well-known mechanisms of biofilm forma-tion belong to proteins and polysaccharides (Kristensen et al2008). There are number of methods that are used to controlbiofilm formation in microbial cells, one of which is thebiochemical technique which controls the structure and archi-tecture of biofilm by modification to diminish biofilm forma-tion; this is usually done by adding enzymes to destroy thestructure of EPS or protein. Although this method is of lowtoxicity and efficiency, it is rendered impractical due to thehigh cost associated with enzyme production (Richards andCloete 2010). Another biochemical method is the use ofsignaling molecules which are responsible for cell–cellcommunication in different bacterial and fungal cultures(Al-Juboori and Yusaf 2012). One of the most famous signal-ing molecules in fungi is farnesol; this aliphatic alcohol blocksthe yeast-to-mycelium conversion (Nickerson et al. 2006).Ethanol, aliphatic alcohol, was used in this study as ananalogue to farnesol. Ethanol exerts different effects whenadded to microbial cultures: it has been stated that it affects

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Fig. 5 Changes in lipid peroxidation (TBARS) and mycelial weight inPenicillium purpurogenum grown in cultures amended with differentethanol concentrations

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Fig. 6 Color changes as measured at 492 nm in Penicilliumpurpurogenum grown in cultures amended with different ethanolconcentrations

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membrane fluidity (Da Silveria et al. 2003) and enhancesproteases production (Meza et al. 2007). Ethanol enters themetabolic network through the gluconeogenic pathway(Mogensen et al. 2006), and hence does not affect the culturemedia or leave unwanted toxic by-products. The results clear-ly show that ethanol exerted a stress-inducing effect on thefungus Penicillium purpurogenum; this was clear from thecatalase production and lipid peroxidation induced at differentethanol concentrations. In addition, the prominent red color ofthe culture decreased as the concentration of ethanol in-creased, which is another indication that ethanol caused stressto the fungus and that the pigment was used to overcome thisstress. This result is in accordance with Palanisami andLakshmanan (2010), who stated that some pigments havebeen reported to possess an antioxidant activity; their decreaseupon stress is attributed to their involvement as antioxidants toprotect the cells, hence their decrease. On the other hand, Hanet al (2005) contradicted this statement and stated that carot-enoid yield increased in the presence of oxidative stress.Fungal pigments usually fall into one of four categories: theshikimate-, terpenoid-, polyketide- and nitrogen-containingpigments (Velsek and Cejpek 2011). Since Penicillium

purpruogenum produces red pigment which falls into thecategory of polyketide pigments (Mendez et al. 2011), whichare either ketides or fatty acids, the first under goes cycliza-tion, while the second undergoes reduction of the carbonylgroups (Velsek and Cejpek 2011), while another possibleexplanation for the decrease in pigment production as theconcentration of ethanol increased is the interference of etha-nol in the first step of pigment cyclization and/or reduction,according to its precise nature. Pigment production is sensitiveto many environmental factors such as light and growth(Velmurugan et al 2010). The addition of ethanol had agrowth-inhibiting effect at higher concentrations; this resultis in accordance with Meza et al. (2007), who stated thatadding ethanol to the fungal culture medium had an adverseeffect on the fungal growth. Due to all these findings, lowethanol concentration was used to study its effect on biofilmformation. The results clearly show that it was the surface-bound protein that was affected by the ethanol added to theculture medium and not EPS which is responsible for cell

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Fig. 7 Surface-bound EPS andsurface-bound protein inPenicillium purpurogenumgrownin cultures amendedwith differentethanol concentrations

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Fig. 8 Cell surface charge (CSC) for Penicillium purpurogenum grownin cultures amended with 2.5 and 5 % v/v ethanol

Ethanol as an external stimuli

Cell wall changes

Catalase

Lipid peroxidation

Mycelial growth

Pigment

Cell surface hydrophobicity

Physiological and Stress response

Surface bound proteins

Exopolysaccharides

Fig. 9 Representation of the changes exerted by ethanol on the fungusPenicillium purpurogenum

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adhesion; this result is in agreement with Meza et al, (2007).The oxidative stress resulting from the addition of alcohol tothe fungus is one of many complicated intra-species commu-nication, the sender represented by the ethanol and the receiv-er represented by morphological changes and oxidativestress (Cottier and Mühlschlegel 2012). Adhesive prop-erties in fungi are expressed by a group of cell-srrfaceproteins called adhesins (Linder and Gustafsson 2008), andthese adhesins could be the main reason biofilm formationtakes place, and, consequently, their depletion by ethanolprohibited biofilm formation. The cell surface charge is oneof the parameters which are used to evaluate cell adhesion tosurfaces (Castellanos et al 1997). The addition of ethanol toPenicillium purpurogenum spores resulted in an increase inthe cell surface charge; this, too, is an indication that surface-bound proteins are the ones involved in cell adhesion, hencebiofilm formation, mainly due to the changes in the net surfacecharge. A representation of the changes which took place afterethanol was added to the fungus is shown in Fig. 9.

In conclusion, fungal adhesion could bemanipulated by theaddition of ethanol which could affect the adhesion of bothcell-to-cell and cell-to-substrate. This low-cost by-productwill offer a safe alternative to existing biofilm and biofoulingcontrol agents, and will also not exert any toxic effects on theenvironment, as it is metabolized by the fungus.

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