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  1. 1. 1 23 Annals of Microbiology ISSN 1590-4261 Ann Microbiol DOI 10.1007/s13213-013-0788-5 The role of ethanol in preventing biofilm formation of Penicillium purpurogenum Sherif M. Husseiny, Hussein Abd El Kareem, Ola M. Gomaa & Riham Talaat
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  3. 3. ORIGINAL ARTICLE The role of ethanol in preventing biofilm formation of 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 no cell 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 and cell–surface adhesion through manipulating cell surface prop- erties. Scanning electron microscopy indicated that the type of surface and its treatment with ethanol controls the adhesion and biofilm formation of Penicillium purpurogenum. Gamma irradiation slightly affected the wettability of polystyrene strips 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 decrease in surface-bound proteins from 0.348 to 0.133 mg/ml, while surface exopolysaccharides showed a minimal decrease. Ethanol induced oxidative stress which reached its peak when 2.5 % v/v ethanol was added to the media; this was represent- ed by both intracellular and extracellular catalase and lipid peroxidation. On the other hand, fungal biomass and pigment showed a decrease as the ethanol concentrations increased. Therefore, ethanol could be employed to control the surface properties of a fungus, and to inhibit biofilm formation to obtain a high surface area for the fungus to be employed in any biotechnological process. Keywords Penicillium purpurogenum . Adhesion . Biofilm formation . Ethanol . Oxidative stress response Introduction The adherence of microbial cells onto surfaces often results in a build-up of aggregates and the formation of what is known as “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 substance which offer protection against biocides and toxins (Stoodley et al. 2004). However, as crucial as it is for microorganisms to form biofilms to protect their integrity and continue their survival in any given harsh environment, biofilm formation is considered a common threat in fields such as the food industry (Simões et al. 2010) and the biomedical field (Hao et al. 2012). It is also perceived as a threat in wastewater treatment reactors for their ability to cause corrosion, odor, and hydrogen sulfide (Jiang and Yuan 2013). Biofilms may act as a harbor for pathogenic microbial cells in drinking water reservoirs (Piriou et al 1997). Problems arising from biofilm formation is due to the cost associated with the losses it causes: the deterioration in plant performance, the decrease in the quality and quantity of the product, the damage of the constructing material, and the cost of cleaning processes or cost of addition al biocides or labor used to replace or clean the tanks (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 formation such 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 controlling biofilm formation include membrane surface modification or biochemical techniques which involve degrading the EPS using enzymes, bacteriophages, and signaling proteins (Al-Juboori and Yusaf 2012). The attachment process between fungal spores and/or hyphae and substrates is considered a very complex process; it mainly depends on the physicochem- ical surface interaction; specific molecular factors being S. M. Husseiny Botany Department, Girl’s College, Ain Shams University, Cairo, Egypt H. A. El Kareem :O. M. Gomaa (*) :R. Talaat Microbiology Department, National Center for Radiation Research and Technology (NCRRT), Cairo, Egypt e-mail: ola_gomaa@hotmail.com Ann Microbiol DOI 10.1007/s13213-013-0788-5 Author's personal copy
  4. 4. glycoproteins, hydrophobins, carbohydrates, and lipids (Priegnitz et al 2012). Biofilm removal or prevention is considered somewhat easier when there is no need for live microbial cells, but it is very difficult to prevent its formation and still keep the cells live and intact. Ethanol is an agro-industrial waste that is produced in huge quantities in Egypt in the process of making sugar from sugar cane; therefore, it is very cheap and abun- dant. It is an aliphatic alcohol that has been used in various ways, when added at 70 % (w/v) it is employed as a disinfec- tant, while at 5–10 % it is bacteriostatic (Sissons et al 1996). It can be used as a carbon source for fungi (Mogensen et al 2006), as a supplemental electron donor to stimulate microbial reduction of nickel and iron (Akob et al 2008), and as an inducer for laccase (phenol oxidase) enzyme in white rot fungi (Alves et al 2004). The fungus Penicillium purpurogenumis a filamentous 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 it has also been characterized as phenol oxidase producer in our laboratory (data unpublished). In the following work, ethanol will be employed to control the adhesion and biofilm forma- tion of Penicillium purpurogenum, through examining some of the parameters controlling microbial adhesion. Materials and methods Fungal isolation and cultivation conditions The fungus used was isolated from soil about 40 km outside Cairo, Egypt. About 10 g were added to 90 ml sterile saline solution and shaken for 1 h. After serial dilution, 0.1 ml of the appropriate 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 was composed 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, and CuSO4·5H2O 0.05 g. After 7 days incubation, the samples were purified by streaking onto clean CZYA plates. Periodical subculturing of fungi was performed on agar slants and stored at 4 °C. The preliminary identification of the isolates was done on water agar plates based on their morphology according to Pitt and Hocking (1985). Morphological study Using CZYA plates with and without ethanol, a glass cover slide was placed at an angle of 45°, the fungus was inoculated at the base of the cover slide, and was left to incubate for 7 days. The cover slides with the grown fungus on the edge were used for scanning electron microscopy as described below. Adhesion and biofilm formation Polystyrene sheet, tin sheet, and glass cover slides were all cut into 0.25×0.5 cm strips. About 100 μl of spore suspension was added to the wells of a round-bottom 96-well microtiter plate, and the wells were divided to groups, each containing strips of polystyrene, tin, or glass, with and without ethanol. Another group was used with ethanol-immersed strips. The plates were left to incubate for 24 h as previously described. The strips were taken from the wells with a sterile forceps and left to dry in air. Scanning electron micrographs of the adhe- sion on the strips was carried out using a JOEL JMS 5600 scanning 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 gold under reduced pressure. The images were captured at magni- fications of ×750 using an electron beam high voltage of 30 kV. Gamma radiation Polystyrene strips were placed each in separate pouches and were used for gamma radiation experiments. Gamma irradia- tion was performed in triplicates at the cobalt source located at NCRRT, 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 polymer did 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-old culture was taken used the broad side of a sterile tip was used to inoculate a set of cultures. Ethanol was added under sterile - conditions to different CZY liquid cultures in 100-ml Erlenmeyer flasks with 20 ml working volume on the day of inoculation to obtain final concentrations of 0, 2.3, 5, 7.5, and 10 % v/v. The Erlenmeyer flasks were incubated under static conditions at 30 °C for 7 days. The cultures were used for the following biochemical assays. Catalase Catalase was measured according to the method of Beers and Sizer (1952). The disappearance of peroxide was followed spectrophotometrically at 240 nm using a Schimadzu UV 2100 spectrophotometer. One Unit was defined as the quantity of catalase that decomposes 1 µmol of H2O2 per min at 25 °C (pH 7.0). The reaction mixture consisted of 0.05 M potassium Ann Microbiol Author's personal copy
  5. 5. phosphate buffer (pH 7) containing 0.059 M hydrogen peroxide. Lipid peroxidation Lipid peroxidation was calculated as the concentration of malondialehyde (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 to Yoshika et al. (1979). Mycelial weight The fungal biomass obtained at the end of incubation period was washed with distilled water and dried in an oven at 70 °C for 24 h. Dry biomass was determined as dry weight per volume. Red pigment assay Red pigment assay was performed using the extracellu- lar fluid (ECF) for each culture; the ECF were used for visible spectrophotometric analysis at 492 nm to test the changes in color for all tested ethanol concentrations (Mendez et al 2011). Exopolysaccharides (EPS) Cultures in the previous experiment were centrifuged at 5,000 rpm for 15 min, the supernatant was removed and 95 % ethanol was added to the cells and incubated at 4 °C overnight 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-sulfuric method (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 a modified method of Castellanos et al. (1997), the cells were harvested, washed twice with PBS, and the pellets resuspend- ed in 10 ml 6 M urea for 90 min at 22 °C. The cell suspension was centrifuged at 1,600 g for 10 min at 10 °C, and the supernatant was used to detect the protein content using Lowry’s method (Lowry et al. 1951) using bovine serum albumin (BSA) as a standard. Cell surface charge Spore suspension of Penicillium purpurogenum was used to detect the cell surface charge in the presence (2.5 and 5 % v/v) and absence of ethanol using the two-phase partitioning assay as described by Castellanos et al (1997). Each system was done separately; the phases 1 and 2 were added consecutively. System I consisted of 7.13 % polyethylene glycol (PEG) in 150 mM NaCl as phase 1 and 8.75 % dextran in 150 mM NaCl as phase 2, while system II consisted of 7.13 % PEG in 150 mM as phase 1 and 8.35 % dextran and 0.4 % dextran sulfate in 150 mM NaCl as phase 2. The results were expressed as Δ log G which is defined by the following equation: 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 the system. Values larger than zero indicate negatively charged cell surface. Control Penicillium purpurogenum culture Ethanol amended Penicillium purpurogenum culture Fig. 1 Scanning electron micrographs of ethanol amended Penicillium purpurogenum cultures as compared to control cultures, magnification ×2000 Ann Microbiol Author's personal copy
  6. 6. Results Morhological changes Cultures of Penicillium purpurogenum were examined for morphological changes after growth in ethanol and in control cultures. The pictures in Fig. 1 clearly show that ethanol-amended cultures exhibited la oose myce- lial network as compared to a tight network in control cultures. To study the effect of ethanol on biofilm formation on different substrates, glass, polystyrene, and tin foil strips were used, each categorized to control and ethanol-amended groups. Figure 2 shows that the different cultures grown in ethanol showed obviously less adhesion on different media as compared to those grown in ethanol-free media. The least growth was exhibited on glass strips, followed by polystyrene strips, 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 Penicillium purpurogenum grown on glass (1, 2), polystyrene strips (3, 4) and tin foil strips (5, 6) in control and ethanol amended microtitre plates, respectively Ann Microbiol Author's personal copy
  7. 7. 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 0.3 and 0.5 kGy and was maximal at 1 kGy (Fig. 3). Biological changes The results in Fig. 4 clearly show that the addition of ethanol to the culture media results in a parallel increase in both extracellular and intracellular catalase; the former is produced in quantity. Catalase activity reached its peak at 6.49 U/ml when 2.5 % v/v was added to the media, after which the activity showed a gradual decrease and reached its minimum of 2.49 U/ml when 10 % v/v ethanol was added to the media. Although intracellular catalase followed the same pattern, the values were below those shown for extracellular catalase. Another indication of stress by ethanol is shown in Fig. 5 which represents the degree of lipid peroxidation and mycelial growth in the presence of different concentrations of ethanol. The figure shows that lipid peroxidation reached its maximum of 0.23 mg/mg mycelia when 2.5 % v/v ethanol was added to the culture medium, above which there was a drop in lipid peroxidation. On the other hand, mycelial growth was main- tained in the presence of 2.5 and 5 % v/v ethanol along with control cultures and was represented as 100 % mycelial growth, above which there was a sharp decrease in mycelial growth that reached only 5 %. Penicillium purpurogenum produces a red pigment, which was affected by the addition of ethanol to the culture medium. The deep rich red color of control cultures showed absorbance of 5.8 but exhibited a lighter shade as the ethanol concentra- tion increased, until it reached an absorbance of 0.97 when 10 % 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 radiation on adhesion of Penicillium purpurogenum on polystyrene strips grown in microtiter plates 0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5 5.5 6 6.5 7 7.5 0 1 2 3 4 5 6 7 8 9 10 11 12 Ethanol (% v/v) Catalase(U/ml) Intracellular CAT Extracellular CAT Fig. 4 Intracellular and extracellular catalase in Penicillium purpurogenum grown in cultures amended with different ethanol concentrations Ann Microbiol Author's personal copy
  8. 8. 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 at different ethanol concentrations, the results varying from 10.3 to 9.5 mg/ml. On the other hand, the surface-bound proteins exhibited a drop which reached 0.133 mg/ml as compared to 0.348 mg/ml in control cultures which means there was about 2.6-fold drop in surface-bound proteins when ethanol was added to the culture medium. To study the degree of cell attachment in the presence of ethanol, the cell surface charge (CSC) was calculated in the presence and absence of ethanol. Figure 8 shows an increase in CSC value from 0.043 in control cultures, to 0.18 for 2.5 % v/v ethanol, and reached a maximum value of 0.32 when 5 % v/v ethanol was added to the culture media. Discussion Biofilm formation is considered a complex process; it is initiated by the adhesion of cells to cells or cells to surface. The adhesion process is comprised of two parts: an initial physical attachment takes place first; this is due to weak, reversible van der Waals forces. Later on, an irreversible strong 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, scanning electron microscopy was used as a monitoring technique (Al-Juboori and Yusaf 2012). The results show that fungal morphology in the presence of ethanol was markedly affected; control cultures showed tightly bound mycelial growth while ethanol-amended cultures showed a loose bound mycelial growth. This result is another confirmation that the presence of non-growth-inhibiting ethanol in the medium had an effect on the surface characteristics and architecture of the growing mycelium. For detection of fungal adhesion on dif- ferent substrates, scanning electron microscopy was also used to monitor the adhesion on glass, polystyrene, and tin foil in the presence and absence of ethanol. The results show that fungi showed less adhesion in the presence of ethanol for all three substrates used; however, there was also a variation in the adhesion even when ethanol was added, the least being on glass, followed by polystyrene, and the highest adhesion being on tin foil. Glass is stated to be a wettable surface while polystyrene is a less wettable surface (Amiri et al 2005). This suggests that the adhesion is related to both the changes on the cell surface and the hydrophobicity of the substrate. This line of evidence is further supported by the fact that using gamma radiation to vary the wettability of polystyrene sur- faces caused an alteration in the adhesion; the same took place when polystyrene sheets were immersed in etha- nol. The use of radiation to decrease the adhesion of a wettable substrate has been used before: UV radiation was used to decrease polystyrene wettability and hence decrease the 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 al 2008). There are number of methods that are used to control biofilm formation in microbial cells, one of which is the biochemical 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 the structure of EPS or protein. Although this method is of low toxicity and efficiency, it is rendered impractical due to the high cost associated with enzyme production (Richards and Cloete 2010). Another biochemical method is the use of signaling molecules which are responsible for cell–cell communication 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 blocks the yeast-to-mycelium conversion (Nickerson et al. 2006). Ethanol, aliphatic alcohol, was used in this study as an analogue to farnesol. Ethanol exerts different effects when added to microbial cultures: it has been stated that it affects 0 0.05 0.1 0.15 0.2 0.25 0 1 2 3 4 5 6 7 8 9 10 11 Ethanol (%v/v) TBARS(mg/mgmycelia) 0 20 40 60 80 100 120 Mycelialweight(%) TBARS Mycelial weight Fig. 5 Changes in lipid peroxidation (TBARS) and mycelial weight in Penicillium purpurogenum grown in cultures amended with different ethanol concentrations 0 1 2 3 4 5 6 7 0 1 2 3 4 5 6 7 8 9 10 11 Ethanol (% v/v) Absorbance Fig. 6 Color changes as measured at 492 nm in Penicillium purpurogenum grown in cultures amended with different ethanol concentrations Ann Microbiol Author's personal copy
  9. 9. membrane fluidity (Da Silveria et al. 2003) and enhances proteases production (Meza et al. 2007). Ethanol enters the metabolic network through the gluconeogenic pathway (Mogensen et al. 2006), and hence does not affect the culture media or leave unwanted toxic by-products. The results clear- ly show that ethanol exerted a stress-inducing effect on the fungus Penicillium purpurogenum; this was clear from the catalase production and lipid peroxidation induced at different ethanol concentrations. In addition, the prominent red color of the culture decreased as the concentration of ethanol in- creased, which is another indication that ethanol caused stress to the fungus and that the pigment was used to overcome this stress. This result is in accordance with Palanisami and Lakshmanan (2010), who stated that some pigments have been reported to possess an antioxidant activity; their decrease upon stress is attributed to their involvement as antioxidants to protect the cells, hence their decrease. On the other hand, Han et 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: the shikimate-, terpenoid-, polyketide- and nitrogen-containing pigments (Velsek and Cejpek 2011). Since Penicillium purpruogenum produces red pigment which falls into the category of polyketide pigments (Mendez et al. 2011), which are either ketides or fatty acids, the first under goes cycliza- tion, while the second undergoes reduction of the carbonyl groups (Velsek and Cejpek 2011), while another possible explanation for the decrease in pigment production as the concentration 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 sensitive to many environmental factors such as light and growth (Velmurugan et al 2010). The addition of ethanol had a growth-inhibiting effect at higher concentrations; this result is in accordance with Meza et al. (2007), who stated that adding ethanol to the fungal culture medium had an adverse effect on the fungal growth. Due to all these findings, low ethanol concentration was used to study its effect on biofilm formation. The results clearly show that it was the surface- bound protein that was affected by the ethanol added to the culture medium and not EPS which is responsible for cell 5 6 7 8 9 10 11 0 1 2 3 4 5 6 7 8 9 10 11 Ethanol (% v/v) SurfaceboundEPS(mg/ml) 0 0.05 0.1 0.15 0.2 0.25 0.3 0.35 0.4 Surfaceboundprotein(mg/ml) Surface bound EPS Surface bound protein Fig. 7 Surface-bound EPS and surface-bound protein in Penicillium purpurogenumgrown in cultures amended with different ethanol concentrations 0 0.05 0.1 0.15 0.2 0.25 0.3 0.35 0 1 2 3 4 5 6 Ethanol (%v/v) CSC Fig. 8 Cell surface charge (CSC) for Penicillium purpurogenum grown in 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 fungus Penicillium purpurogenum Ann Microbiol Author's personal copy
  10. 10. adhesion; this result is in agreement with Meza et al, (2007). The oxidative stress resulting from the addition of alcohol to the 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 oxidative stress (Cottier and Mühlschlegel 2012). Adhesive prop- erties in fungi are expressed by a group of cell-srrface proteins called adhesins (Linder and Gustafsson 2008), and these adhesins could be the main reason biofilm formation takes place, and, consequently, their depletion by ethanol prohibited biofilm formation. The cell surface charge is one of the parameters which are used to evaluate cell adhesion to surfaces (Castellanos et al 1997). The addition of ethanol to Penicillium purpurogenum spores resulted in an increase in the cell surface charge; this, too, is an indication that surface- bound proteins are the ones involved in cell adhesion, hence biofilm formation, mainly due to the changes in the net surface charge. A representation of the changes which took place after ethanol was added to the fungus is shown in Fig. 9. In conclusion, fungal adhesion could be manipulated by the addition of ethanol which could affect the adhesion of both cell-to-cell and cell-to-substrate. This low-cost by-product will offer a safe alternative to existing biofilm and biofouling control agents, and will also not exert any toxic effects on the environment, as it is metabolized by the fungus. 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