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

2. 1 23
Your article is protected by copyright and
all rights are held exclusively by Springer-
Verlag Berlin Heidelberg and the University
of Milan. This e-offprint is for personal
use only and shall not be self-archived in
electronic repositories. If you wish to self-
archive your article, please use the accepted
manuscript version for posting on your own
website. You may further deposit the accepted
manuscript version in any repository,
provided it is only made publicly available 12
months after official publication or later and
provided acknowledgement is given to the
original source of publication and a link is
inserted to the published article on Springer's
website. The link must be accompanied by
the following text: "The final publication is
available at link.springer.com”.

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. 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. 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. 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. 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. 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. 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. 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.
References
Akob DM, Mills HJ, Gihring TM, Kerkhof L, Stucki JW, Anastacio AS,
Chin KJ, Kusel K, Palumbo AV, Watson DB, Kostka JE (2008)
Functional diversity and electron donor dependence of microbial
populations capable of U(VI) reduction in radionuclide-
contaminated subsurface sediments. Appl Environ Microbiol 74:
3159–3170
Al-Juboori RA, Yusaf T (2012) Biofouling in RO system: Mechanisms,
monitoring and controlling. Desalination 302:1–23
Alves AMCR, Record E, Lomascolo A, Scholtmeijer K, Asther M,
Wessels JGH, Wosten HAB (2004) Highly efficient production of
laccase by the basidiomycete Pycnoporus cinnabarinus. Appl
Environ Microbiol 70:6379–6384
Amiri A, Cholodowski D, Bompeix G (2005) Adhesion and germination
of waterborne and airborne conidia of Penicillium expansum to
apple and inert surfaces. Physiol Mol Plant Pathol 67:40–48
Beers RF, Sizer IW (1952) A spectrophotometric method for measuring
the breakdown of hydrogen peroxide by catalase. J Biol Chem 195:
130–140
Castellanos T, Ascencio F, Bashan Y (1997) Cell-surface hydrophobicity
and cell-surface charge of Azospirillum spp. FEMS Microbiol Ecol
24:159–172
Chaplin MF, Kennedy JF (1986) Carbohydrate analysis: A practical
Approach. Oxford University Press, Oxford, pp 1–2
Cottier F, Mühlschlegel FA (2012) Communication in fungi. Int J
Microbiol. doi:10.1155/2012/351832
Da Silveria MG, Golovina EA, Hoekstra FA, Rombouts FM, Abee T
(2003) Membrane fluidity adjustments in ethanol-stressed
Oenococcus oeni cells. Appl Environ Microbiol 69:5826–5832
Ferrnandez-Olmos A, Garcia-Castillos M, Maiz L, Lamas A, Baquero F,
Canton R (2012) In vitro prevention of Pseudomonas aeruginosa
early biofilm formation with antibiotics used in cystic fibrosis pa-
tients. Int J Antimicrob Agents 40:173–176
Han R, Zhao WJ, Gao YY, Yuan JM (2005) effect of oxidative stress and
exogenous β-carotene on sclerotial differentiation and carotenoid
yield of Penicillium sp. PT95. Lett Appl Microbiol 40:412–417
Hao C, Zhou Z, Peng C, Gen WUX, Ge Z (2012) Inhibitory effect of
extracellular polysaccharides EPS-II from Pseudoalteromonas on
Candida adhesion to cornea in vitro. Biomed Environ Sci 25:210–
215
Jiang G, Yuan Z (2013) Synergistic inactivation of anerobic wastewater
biofilm by free nitrous acid and hydrogen peroxide. J Hazard Mater
250–252:91–98
Kristensen JB, Meyer RL, Laursen BS, Shipovskov S, Besenbacher F,
Poulsen CH (2008) Antifouling enzymes and the biochemistry of
marine settlement. Biotechnol Adv 26:471–481
Linder T, Gustafsson CM (2008) Molecular phylogenetics of
ascomycotal adhesins-A novel family of putative cell-surface adhe-
sive proteins in fission yeasts. Fungal Genet Biol 45:485–497
Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein mea-
surement with Folin phenol reagent. J Biol Chem 193:256–275
Mendez A, Perez C, Montanez JC, Martinez G, Aguilar CN (2011) Red
pigment production by Penicillium purpurogenum GH2 is influ-
enced by pH and temperature. J Biomed Biotechnol 12:961–968
Meza JC, Auria R, Lomascolo A, Sigoillot JC, Casalot L (2007)
Role of ethanol on growth, laccase production and protease
activity in Pycnoporus cinnabarinus ss3. Enzyme Microb Technol
41:162–168
Mogensen J, Nielsen HB, Hofmann G, Nielsen J (2006) Transcription
analysis using high-density micro-arrays of Aspergillus nidulans
wild-type and creA mutant during growth on glucose or ethanol.
Fungal Genet Biol 43:593–603
Monteiro AS, Miranda TT, Lula I, Dendai A, Sinisterra RD, Santoro MM,
Santos VL (2011) Inhibition of Candida albicans CC biofilms
formation in polystyrene plate surfaces by biosurfactant produced
by Trichosporon montevideense CLOA72. Colloids Surf B 84:467–
476
Nehad EA, El-Shamy AR (2010) Physiological studies on the production
of exopolysachharides by fungi. Agric Biol J N Am 1:1303–1308
Nickerson KW, Atkin AL, Horny JM (2006) Quorum sensing in dimor-
phic fungi: farnesol and beyond. Appl Environ Microbiol 72:3805–
3813
Palanisami S, Lakshmanan U (2010) Role of copper in poly R-478
decolorization by the marine cyanobacterium Phormidium
valderianum, BDU140441. World J Microbiol Biotechnol 27:669–
677
Piriou P, Dukan S, Levi Y, Jarrige PA (1997) Prevention of bacterial
growth in drinking water distribution systems.Water Sci Technol 35:
283–287
Pitt JI, Hocking AD (1985) Fungi and Food Spoilage. Academic, Sydney
Priegnitz BE, Wargenau A, Brandt U, Rohde M, Dietrich S, Kwade A,
Krull R, Fleißner (2012) The role of initial spore adhesion in pellet
and biofilm formation in Aspergillus niger. Fungal Genet Biol 49:
30–38
Richards M, Cloete TE (2010) Nanoenzymes for biofilm removal. In:
Cloete TE, Dekwaadstenient M, Botes M, Lopez-Romero JM (eds)
Nanotechnology in Water Treatment Applications. Caister
Academic, Norfolk, pp 89–102
Scopel M, Abraham WR, Henriques AT, Macedo AJ (2013) Dipeptide
cis-cyclo(Leucyl-Tyrosyl) produced by sponge associated
Penicillium sp. F37 inhibits biofilm formation of the pathogenic
Staphylococcus epidermis. Bioorganic Med Chem Lett 23:624–626
Simões M, Simões LC, Vieira MJ (2010) A review of current and
emergent biofilm control strategies. LWT-Food Sci Technol
43:573–583
Ann Microbiol
Author's personal copy

11. Sissons CH, Wong L, Cutress TW (1996) Inhibition by ethanol of the
growth of biofilm and dispersed microcosm dental plaques. Arch
Oral Biol 1:27–34
Stoodley LH, Costerton JW, Stoodley P (2004) Bacterial biofilms: from
the natural environment to infectious diseases. Nat Rev Microbiol 2:
95–108
Velmurugan P, Lee YH, Venil CK, Lakshmanaperumalsamy P, Chae JC,
Oh BT (2010) Effect of light on growth, intracellular and
extracellular pigment production by five pigment-producing
filamentous fungi in synthetic medium. J Biosci Bioeng 109:
346–350
Velsek J, Cejpek K (2011) Pigments of higher fungi: A review. Czeck J
Food Sci 29:87–102
Yoshika T, Kawada K, Shimda T, Mori M (1979)Lipid peroxidation in
maternal and cord blood andprotective mechanism against activated
oxygentoxicity within blood. Am J Obstet Gynecol135:372–376
Ann Microbiol
Author's personal copy