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APPLIED MICROBIAL AND CELL PHYSIOLOGY
Effect of growth temperature, surface type and incubation
time on the resistance of Staphylococcus aureus biofilms
to disinfectants
Marwan Abdallah &Gabrielle Chataigne &
Pauline Ferreira-Theret &Corinne Benoliel &
Djamel Drider &Pascal Dhulster &Nour-Eddine Chihib
Received: 8 October 2013 / Revised: 16 December 2013 / Accepted: 17 December 2013 / Published online: 16 January 2014
#Springer-Verlag Berlin Heidelberg 2014
Abstract The goal of this study was to investigate the effect
of the environmental conditions such as the temperature
change, incubation time and surface type on the resistance of
Staphylococcus aureus biofilms to disinfectants. The
antibiofilm assays were performed against biofilms grown at
20 °C, 30 °C and 37 °C, on the stainless steel and polycar-
bonate, during 24 and 48 h. The involvement of the biofilm
matrix and the bacterial membrane fluidity in the resistance of
sessile cells were investigated. Our results show that the
efficiency of disinfectants was dependent on the growth tem-
perature, the surface type and the disinfectant product. The
increase of growth temperature from 20 °C to 37 °C, with an
incubation time of 24 h, increased the resistance of biofilms to
cationic antimicrobials. This change of growth temperature
did not affect the major content of the biofilm matrix, but it
decreased the membrane fluidity of sessile cells through the
increase of the anteiso-C19 relative amount. The increase of
the biofilm resistance to disinfectants, with the rise of the
incubation time, was dependent on both growth temperature
and disinfectant product. The increase of the biofilm age also
promoted increases in the matrix production and the mem-
brane fluidity of sessile cells. The resistance of S. aureus
biofilm seems to depend on the environment of the biofilm
formation and involves both extracellular matrix and mem-
brane fluidity of sessile cells. Our study represents the first
report describing the impact of environmental conditions on
the matrix production, sessile cells membrane fluidity and
resistance of S. aureus biofilms to disinfectants.
Keywords Staphylococcus aureus .Abiotic surfaces .
Environmental conditions .Biofilm resistance .Disinfectants
Introduction
The Gram-positive bacterium Staphylococcus aureus is an
opportunistic human pathogen. This bacterium is not only an
established nosocomial pathogen but also one of the great
common causes of food illness outbreaks (Crago et al. 2012;
Kerouanton et al. 2007). When growing in foods, S. aureus
strains are able to produce staphylococcal enterotoxins which
are the causative agents of staphylococcal food poisonings
(Normanno et al. 2007). It is now established that, in natural
ecosystems, bacteria live attached to surfaces and form a
complex three-dimensional community, called biofilm
(Jenkinson and Lappin-Scott 2001). Moreover, several studies
have shown the ability of S. aureus to attach on food contact
surfaces such as metal, rubber, polypropylene, glass, wood
and food products (Simões et al. 2010). The persistence of
biofilms on these surfaces presents a continuous source of
food contamination resulting in food poisoning. Thus, the
fight against biofilms represents a challenge for the food
sector in order to ensure the production of safety products
(Carpentier and Cerf 1993; Costerton et al. 1999;Simõesetal.
2010).
Biofilm is a community of microorganisms irreversibly
attached to a substratum and embedded in an extracellular
M. Abdallah :G. Chataigne :D. Drider :P. Dhulster :
N.<E. Chihib (*)
Laboratoire de Procédés Biologiques, Génie Enzymatique et
Microbien (ProBioGEM), IUT A/Polytech’Lille, Université de Lille
1-Science et Technologies, Avenue Paul Langevin, 59655 Villeneuve
d’Ascq Cedex, France
e-mail: nour-eddine.chihib@univ-lille1.fr
M. Abdallah :P. Ferreira-Theret:C. Benoliel
Laboratoire SCIENTIS, Parc Biocitech, 102, Avenue Gaston
Roussel, 93230 Romainville, France
Appl Microbiol Biotechnol (2014) 98:2597–2607
DOI 10.1007/s00253-013-5479-4
polymeric matrix that confers resistance for the involved
microorganisms (Donlan 2002). It has been reported that, in
the food industry, factors such as pH, temperature changes,
surface type and nutrient availability are among the relevant
factors that influence the biofilm formation of S. aureus (Da
Silva Meira et al. 2012; Herrera et al. 2007; Marques et al.
2007). These factors may also enhance the resistance of sessile
cells to disinfecting agents (Bae et al. 2012; Belessi et al.
2011; Chavant et al. 2004;DaSilvaMeiraetal.2012;
Joseph et al. 2001; Marques et al. 2007).
Several studies have reported the persistence of pathogenic
bacteria on food and on medical equipments despite the use of
cleaning and disinfection procedures (Brooks and Flint 2008;
Simões et al. 2010; Srinivasan et al. 2003). The persistence of
biofilms is probably due to the failure of disinfectant products,
the inappropriate disinfection protocols and the increased
tolerance to antimicrobial agents (Poulsen 1999). It is now
established that the cells under the biofilm state are more
resistant to disinfectants than their planktonic counterparts.
However, the precise mechanism, by which the biofilms resist
to disinfecting agents, remains unclear. It appears to be a
multifactorial process linked to several parameters such as
the biofilm structure, the extracellular matrix and the physio-
logical state of sessile cells (Bridier et al. 2011a).
Microorganisms are often able to overcome the inhibitory
effect of hostile conditions and respond in a variety of ways, to
change their cell physiology, in an attempt to adapt to new
environmental conditions. An important adaptive response of
bacterial cells to non-optimal growing conditions is the mod-
ifications observed in their membrane lipids (Chihib et al.
2005). The bacterial membranes, composed primarily of phos-
pholipids and proteins, constitutes the first line of bacterial
defense against biocides. Moreover, the phospholipid fatty
acyl chains, which are influenced by the environmental con-
ditions, determine the fluidity of bacterial membranes and
may hinder the penetration of antimicrobial agents (Alvarez-
Ordonez et al. 2008; Dubois-Brissonnet et al. 2011;Toetal.
2002). Hence, the investigation of environmental conditions
effect on the biofilm formation, and resistance to antimicro-
bials, is crucial for the understanding of biofilm resistance
mechanisms and for the development of efficient disinfection
protocols and biocidal agents (Bridier et al. 2011a).
In this regard, our goal is to study the effect of growth
temperature and incubation time on the S. aureus biofilm
formation on stainless steel and polycarbonate, two surfaces
widely encountered in the food equipment. The temperatures
used for this study were 20 °C, 30 °C and 37 °C. Thereafter,
the effect of these environmental factors was investigated on
the biofilm resistance towards disinfectants. In addition, the
extracellular matrix and the shifts in the membrane fatty acid
profile of sessile S. aureus, grown under the conditions afore-
mentioned, were characterized in order to understand the
mechanisms of this resistance.
Material and methods
Bacterial strain and culture conditions
The bacterial strain used for this study was S. aureus CIP 4.83,
stored at −80 °C in Tryptic Soy Broth (TSB; Biokar
Diagnostics, France) containing 40 % (v/v) of glycerol.
Precultures were inoculated by 100 μl from frozen tubes and
grown in 5 ml TSB at the same temperature of culture. The
preculture of 20 °C were incubated for 48 h, while the 30 °C
and 37 °C precultures were incubated for 24 h. The main
cultures were prepared by inoculating 5×10
4
CFU/ml from
the preculture tubes into 50 ml of TSB in sterile 500-ml flasks.
Cultures were incubated under shaking (160 rpm) at 20 °C,
30 °C or 37 °C and cultures were harvested in the late
exponential phase.
Slides preparation
The stainless steel (304 l) (Equinox, France) and polycarbon-
ate slides (Plexilux, France), (2.5×5 × 0.1 cm), were cleaned
by soaking in ethanol 95° (Fluka, Sigma-Aldrich, France) for
an overnight to remove grease and then rinsed in water. Slides
were then soaked in 500 ml of TDF4 detergent (5 %)
(Franklab SA, France), for 20 min at 50 °C under agitation.
The slides were thoroughly rinsed five times, for 1 min
under agitation, in 500 ml of distilled water at 20 °C to
eliminate detergent residues and three times in ultrapure
water (Milli-Q® Academic, Millipore, France). Stainless steel
slides were air-dried and sterilized by autoclaving at 121 °C
for 15 min. Polycarbonate slides were sterilized in ethanol 95°
(10 min).
Cell suspension preparation and biofilm formation assays
S. aureus cells were harvested by centrifugation for 10 min at
3,500× g(20 °C).Bacteria were washed twice with 20 ml
potassium phosphate buffer (PB; 100 mM, pH 7) and finally
resuspended in 20 ml of the same buffer. To disperse cells, a
sonication at 37 kHz was carried out according to Arad et al.
(2013), during 5 min at 25 °C (Elmasonic S60H, Elma,
Germany). Subsequently, bacteria were resuspended in the
PB to a cell concentration of 1.10
8
CFU/ml by adjusting the
optical density to OD
620nm
=0.110±0.005 using UV/visible
light spectrophotometer (Ultrospec 1100 pro; GE Healthcare,
formerly Amersham Biosciences, United Kingdom).
Standardized cell suspensions were diluted 10-fold for the
bacterial adhesion experiments (10
7
CFU/ml). The sterile
slides were placed in the horizontal position in Petri dishes.
The upper face was covered by 3 ml cell suspensions
(10
7
CFU/ml) and incubated at 20 °C for 60 min. Then the
coupons were removed using sterile forceps and gently rinsed
by dipping into 30 ml of PB to remove excess liquid droplets
2598 Appl Microbiol Biotechnol (2014) 98:2597–2607
and loosely attached cells. Thereafter, the upper face was
covered with 2 ml TSB and the coupons were incubated, at
the same temperature of cultures (20 °C, 30 °C or 37 °C), for
an incubation times of 24 or 48 h. For the biofilm grown
during 48 h, the culture medium was changed after 24 h of
growth. After 24 and 48 h, the slides were withdrawn and
rinsed twice by gently dipping into 30 ml of PB to remove
loosely attached cells. These coupons were used for the quan-
tification of bacterial cell concentration, for the antibiofilm
assays and for the quantification of proteins and carbohy-
drates. For the quantification of bacterial biofilm biomasses,
the cells were detached in 30 ml of PB by vortexing during
30 s followed by a sonication during 5 min. Then the tubes
were vortexed again during 30 s and serial dilutions were
realized in Tryptone Salt broth (TS; Biokar Diagnostics,
France). Samples of 100 μl were spread onto Tryptic Soy
Agar broth plates (TSA; Biokar Diagnostics, France) and
incubated at 37 °C for 24 h. After the incubation time, the
number of viable and culturable cells was counted on the
plates and the results are expressed in log CFU/cm
2
. The
results represent the mean of three independent experiments.
Two coupons were studied for each experiment.
Antibiofilm assays
For the antibiofilm treatments, the rinsed coupons were im-
merged vertically in 30 ml of the disinfectant solution and
incubated during 5 or 15 min for the antibacterial assays (as
recommended by the manufacturer; SCIENTIS, France).
Compositions and characteristics of disinfectants are shown
in Table 1. Afterwards, the coupons were drawn from the
disinfectant solution and immersed in a neutralizing solution
to stop the antibacterial action. The neutralizer contains a
combination of Tween 80 (30 g/l), Saponin (30 g/l), Lecithin
(30 g/l), Sodium Thiosulphate (5 g/l), L-Histidin (1 g/l) and TS
broth (9.5 g/l) (Tote et al. 2010). The viable and culturable
cells were detached and counted as described above. For the
control assays, the disinfectant solutions were replaced by TS
broth solution. The results represent the average of three
independent experiments. Two coupons were studied for each
experiment.
Quantification of carbohydrate and protein amounts
in the biofilm matrix
The protein and carbohydrate concentrations in the matrix of
biofilms, grown on stainless steel and polycarbonate, were
determined after 24 and 48 h of incubation. The protein
concentrations were quantified using the Bradford method
with bovine serum albumin as the standard (Bradford 1976).
The carbohydrate quantification was performed with the
phenol-sulfuric method with glucose as the standard (Dubois
et al. 1956). After the biofilms rinsing with the physiological
water, the biofilm was recovered by scraping surface, aspirat-
ing and expelling at least ten times with 6 ml of ultrapure
water. The suspensions were vortexed during 30 s, followed
by a sonication (37 kHz, Elmasonic S60H, Elma, Germany),
in 15 ml conical tubes during 5 min at 25 °C. The cells were
removed by centrifugation at 5,000 × gduring 15 min, the
supernatants were filtered through 0.2-μm Millipore filters
and then used for the biochemical assays. The results were
presented in μg/cm
2
as the mean of three independent exper-
iments and in each experiment two slides were used.
Cellular fatty acids extraction and analysis
S. aureus embedded in biofilm were collected after scrapping
cells from the rinsed coupons and resuspended in 10 ml of
PB. Then the tubes were vortexed for 30 s. Subsequently, the
sonicationwas carried out at37 kHz during 5 min and the cells
were dispersed by vortexing for 30 s. The cells were harvested
by centrifugation (10,000 × g, 10 min at 4 °C). The superna-
tant was discarded and the pellets, containing about 10
9
CFU,
were washed twice with cold distilled water. Then, 1 ml of the
saponification solution was used to resuspend the washed
pellet and transferred to extraction tube. Subsequently, cells
were submitted to the saponification and methylation. Fatty
acid methyl esters extraction was realized as described previ-
ously by Chihib et al. (2003). Methyl esters analysis were
performed on a GC-2014 gas chromatograph (Shimadzu,
Japan) equipped with a Zebron ZB-FFAP (30 m ×0.25 mm)
capillary column (Phenomenex, Australia), and connected to
Thermo-finnigan Trace DSQ mass spectrometer (Thermo
Tabl e 1 Compositions and characteristics of disinfectant products
Sanitizers Antimicrobial Final concentration
(ppm or %)
a
Action time
(min)
a
P1 DMPAP
a
315 15
PHMB 100
P2 DDAC 137.5 15
P3 DDAC 80 15
Perfume
b
150
P4 DDAC 125 5
ADBAC 475
P5 DDAC 490 15
BDA 180
P6 Ethanol 62 % 5
BDA 1,500
ADBAC alkyldimethylbenzylammonium chloride, BDA bis(3-
aminopropyl) dodecylamine, DMPAP N-didecyl-N-methyl-poly(oxyethyl)
ammonium propionate, DDAC didecyldimethylammonium chloride,
PHMB polyhexamethylene biguanide
a
The concentration and the action time were recommended by the man-
ufacture (SCIENTIS)
b
ZESTY 7 PC109964 perfume contains 47.24 % of limonene and 7.71 %
of linalool (Kao Corporation, Spain)
Appl Microbiol Biotechnol (2014) 98:2597–2607 2599
Fisher Scientific, USA). Samples were injected in split mode.
Chromatographic separation was carried out by a temperature
gradient program beginning with a 5-min isothermal step at
70 °C followed by an increase to 250 °C at 4 °C/min. Helium
was used as the carrier gas. The injector and MS interface
were maintained at 260 °C and 280 °C, respectively. Electron
impact mass spectra were recorded at 70 eV. Masses were
acquired in TIC between m/z 50 at 600 when the acceleration
voltage was turned on after a solvent delay of 3 min. All data
were processed by Xcalibur software (Thermo Fisher
Scientific, USA). All compounds were identified by compar-
ing both the MS spectra and retention index with those avail-
able in libraries, i.e., NIST, Wiley. The results represent the
average of three independent experiments and each experi-
ment are done in duplicate.
Data analysis
The results are presented as mean values and the standard
error to the mean (SEM). Data analysis was performed using
Sigma Plot 11.0 (Systat Software, USA), using one-way
ANOVA (Tukey’s method) to determine the significance of
differences.
Results
Effect of growth temperature, incubation time and surface
type on the biofilm formation
The effect of the growth temperature was investigated on the
S. aureus biofilm formation,on stainless steel and polycarbon-
ate, during 24 and 48 h. The biomasses of S. aureus biofilms,
grown at 20 °C, 30 °C and 37 °C, are shown in Table 2.
The results show that the populations of S. aureus,on
stainless steel and polycarbonate after 24 h, were similar
whatever the temperature studied. Under these conditions,
the bacterial density was of ca. 8.1 log CFU/cm
2
(Table 2).
The increase of the incubation time from 24 to 48 h did not
affect significantly the bacterial density of S. aureus biofilms.
The average of bacterial density of biofilms grown at 20 °C,
30 °C and 37 °C during 48 h was of ca. 8.3 log CFU/cm
2
(Table 2).
Effect of growth temperature and surface type on the biofilm
resistance towards disinfectants
The antibiofilm effect of the disinfectants were performed on
the biofilms grown during 24 h at 20 °C, 30 °C and 37 °C, on
stainless steel and polycarbonate. The viable and culturable
populations recovered after the biofilm treatments, with the
TS broth and disinfectants, are shown in Fig. 1. The results
show that the TS, used as negative control, slightly reduced
the initial population of S. aureus biofilms whatever the
growth temperature studied (Fig. 1). The viable and culturable
biomass, recovered after a treatment of 15 min, was ranged
from 7.5 to 8.1 log CFU/cm
2
on both surfaces (Fig. 1). The
bacterial biomass recovered from biofilms exposed to P1, P2,
P3 and P4 increased when the growth temperature of biofilm
increased for both surfaces. When exposed to the P1 product
for 15 min, the biofilms grown on stainless steel at 20 °C,
30 °C and 37 °C showed respectively a viable and culturable
count of 2.6, 6.0 and 6.2 log CFU/cm
2
(Fig. 1). When the
biofilms were grown at 20 °C, 30 °C and 37 °C, the treatment
with P2 product for 15 min resulted in a reduction of the initial
population respectively to 3.3, 3.8 and 4.3 log CFU/cm
2
on
the stainless steel and respectively to 1.8, 5.3 and 5.6 log
CFU/cm
2
on the polycarbonate. In addition, the results
showed that the surface type has a significant effect on the
efficiency of disinfectants and this effect was growth temper-
ature and product dependent. The treatment of biofilms grown
at 20 °C with P3 for 15 min reduced completely the initial
population of biofilms grown on stainless steel, while the
biofilms grown on polycarbonate showed a viable and
culturable count of 3.3 log CFU/cm
2
(Fig. 1). The cultivable
biomass recovered from biofilms grown on polycarbonate at
37 °C, after the treatment with the P3 disinfectant, was higher
than this recovered from biofilms grown on stainless steel.
Figure 1shows that the biomass recovered from biofilm
grown at 37 °C on stainless steel and polycarbonate was
respectively about 5.4 and 7.0 log CFU/cm
2
after the treat-
ment with the P3 product (Fig. 1). However, when the
biofilms grown at 37 °C was exposed to P4 during 5 min,
the viable and culturable biomass was about 5.7 and 4.7 log
CFU/cm
2
, respectively, on stainless steel and polycarbonate
(Fig. 1). The antibiofilm treatment with the P6 product during
5 min resulted in a complete reduction of the initial population
Tabl e 2 Density of Staphylococcus aureus in the biofilms grown on the
stainless steel and polycarbonate at 20 °C, 30 °C and 37 °C for 24 and
48 h
Log CFU/cm
2
Stainless steel Polycarbonate
Incubation
time
24 h 48 h 24 h 48 h
20 °C 8.4 ±0.2
a,x,X
8.2± 0.2
a,x,X
8.0± 0.2
a,x,X
8.0± 0.2
a,x,X
30 °C 8.0 ±0.2
a,x
8.4± 0.1
a,b,x
8.2± 0.2
a,x,X
8.4± 0.1
b,x,X
37 °C 8.2 ±0.2
a,x,X
8.6± 0.2
b,x,X
8.0± 0.2
a,x,X
8.3± 0.3
a,b,x,X
The data represent the mean of recovered viable and culturable cells count
(log CFU/cm
2
) ± SEM. Between growth temperatures (a, b), attachment
surfaces (x) and among different incubation time (X) under the same
condition, the mean values with the same letters are not significantly
different (P>0.05)
2600 Appl Microbiol Biotechnol (2014) 98:2597–2607
of biofilms grown on the polycarbonate whatever the growth
temperature (Fig. 1). However, the biofilms grown at 20 °C,
30 °C and 37 °C on stainless steel showed respectively a
viable and culturable count of 2.0, 3.8 and 2.3 log CFU/cm
2
after the treatment with the P6 product (Fig. 1). The
antibiofilm treatment with P5 product during 15 min resulted
in a complete reduction of viable and cultivable bacterial
populations (0 log CFU/cm
2
), except for the biofilms grown
on stainless steel at 37 °C (Fig. 1). Under this condition, the
antibiofilm treatment with the P5 product resulted in a viable
and culturable count of 2.8 log CFU/cm
2
.
Effect of the incubation time, surface type and growth
temperature on the biofilm resistance
This investigation was carried out only with the P2, P5 and P6
disinfectants. The results of Table 3show that the treatment of
biofilms grown during 48 h with TS (negative control) slightly
reduced the initial population which was ranged from 7.8 to
8.3 log CFU/cm
2
on stainless steel and polycarbonate
(Table 3). Our findings (Table 3) indicated that the increase
of the incubation time to 48 h affected the efficiency of
disinfectants. The biomass recovered from the biofilm grown
on stainless steel at 20 °C, after the antibiofilm treatment with
the P2 product, increased from 3.3 to 5.0 log CFU/cm
2
(P<0.05) when the incubation time increased from 24 to
48 h (Fig. 1and Table 3). Similar results were obtained when
the biofilms were grown on the polycarbonate at 20 °C (Fig. 1
and Table 3). The increase of incubation time at 30 °C and
37 °C did not show any significant effect on the bacterial
density recovered from biofilms when exposed to the P2
product whatever the surface used (P>0.05) (Fig. 1and
Tab le 3). In addition, the increase of growth temperature from
20 to 37 °C, with an incubation time of 48 h, increased the
sensitivity of biofilms to P2 product. The bacterial biomass
recovered, after P2 treatment, decreased from 5.0 to 3.6 log
CFU/cm
2
(P<0.05) and from 6.4 to 5.2 log CFU/cm
2
(P<0.05), respectively, on the stainless steel and polycarbon-
ate when the growth temperature increased from 20 °C to
37 °C (Table 3).
Tab le 3also shows that the P5 product resulted in a com-
plete reduction of the initial population (0 log CFU/cm
2
),
whatever the surface type, when biofilms were grown at
20 °C during 48 h. However, the increase of the incubation
time from 24 to 48 h at 30 °C resulted in a drastic decrease of
biofilms sensitivity to the P5 product. The biomass recovered,
Fig. 1 Effect of disinfectants on Staphylococcus aureus biofilms grown
during 24 h, on stainless steel (a) and polycarbonate (b), at 20 °C (light
gray column), 30 °C (dark gray column) and 37 °C (black column). Data
represent the mean of recovered viable and culturable cells count (log
CFU/cm
2
) ± SEM.
‡
Product not recommended by the manufacture for the
polycarbonate surface
Tabl e 3 Effect of disinfectants on Staphylococcus aureus biofilms grown during 48 h, on stainless steel (SS) and polycarbonate (PC) substrates, at
20 °C, 30 °C and 37 °C
Log CFU/cm
2
20 °C 30 °C 37 °C
SS PC SS PC SS PC
TS 7.8± 0.5
a,x
7.9± 0.3
a,x
8.2± 0.2
a,x
8.1± 0.3
a,x
8.3± 0.4
a,x
7.9± 0.5
a,x
P2 5.0± 0.1
a
6.4± 0.3
a
4.7± 1.1
a,b,x
6.4± 0.5
a,x
3.6± 0.8
b,X
5.2± 0.1
X
P5 0.0± 0.0
x,X
0.0± 0.0
x,X
6.5± 0.4 5.4± 0.2 2.9± 0.2
X
1.6± 0.7
P6 0.0± 0.0
a,x,X
0.0± 0.0
a,x,X
0.0± 0.0
a,x
0.0± 0.0
a,x,X
0.0± 0.0
a,x
0.0± 0.0
a,x,X
The data represent the mean of recovered viable and culturable cells count (log CFU/cm
2
) ± SEM. Between growth temperatures (a, b), attachment
surfaces (x) and among different incubation time (X) under the same condition, the mean values with the same letters are not significantly different
(P>0.05)
Appl Microbiol Biotechnol (2014) 98:2597–2607 2601
after the treatment of biofilms grown at 30 °C for 48 h with P5,
was 6.5 and 5.4 log CFU/cm
2
, respectively, on stainless steel
and polycarbonate (P<0.05) (Table 3). However, no signifi-
cant difference was observed in the P5 efficiency, against the
biofilms grown at 37 °C, with the increase of the incubation
time on both stainless steel and polycarbonate (P>0.05). The
recovered bacterial density was 2.9 and 1.6 log CFU/cm
2
,
respectively, on stainless steel and polycarbonate after the
treatment with the P5 disinfectant (Table 3). The results of
P6 product treatments indicated that this disinfectant reduced
the viable count to 0 log CFU/cm
2
whatever the temperature
used (Table 3).
Effect of growth temperature, incubation time and surface
type on the extracellular matrix production
The effect of the growth temperature and the incubation time
was investigated on the major biofilm matrix components.
The aim of this investigation was to understand the involve-
ment of the biofilm matrix in the resistance to disinfecting
agents. The results of Table 4show that the protein and
carbohydrate levels in the biofilm matrix were dependent on
the surface type, time and temperature of incubation. When
the biofilms were grown on stainless steel during 24 h, no
significant effect was observed on the proteins concentration
with the increase of the growth temperature (Table 4). The
average concentration of proteins was about 11.2 μg/cm
2
in
the matrix of biofilms grown for 24 h on stainless steel
whatever the growth temperature used (Table 4). However,
when the biofilms were grown on the polycarbonate during
24 h, the proteins concentration increased from 10.7 to
17.9 μg/cm
2
(P<0.05) and from 10.7 to 14.4 μg/cm
2
(P>0.05) when the temperature was raised from 20 °C to
30 °C and from 20 to 37 °C, respectively (Table 4).
Increasing the incubation time from 24 to 48 h resulted in a
significant increase of the total proteins concentration what-
ever the growth temperature used (P<0.05), except for the
biofilm grown at 30 °C on polycarbonate (P>0.05) (Table 4).
When the biofilms were grown at 20 °C, 30 °C and 37 °C, the
protein concentration significantly increased 1.7-, 1.8- and
2.0-fold, respectively, on stainless steel (P<0.05), and in-
creased 1.9-, 1.1- and 2.1-fold, respectively, on polycarbonate
(Table 4).
The carbohydrate amounts in the 24-h biofilm matrix were
similar whatever the surface type and the temperature of
incubation (Table 4). The average of the total carbohydrates
concentration was of ca. 5.3 and 6.3 μg/cm
2
in the matrix of
biofilms grown on stainless steel and polycarbonate, respec-
tively (Table 4). However, the increase of incubation time to
48 h significantly increased the concentration of carbohy-
drates (P<0.05), except for the biofilms cultivated on stainless
steel at 20 °C (P>0.05). Table 4also shows that the carbohy-
drates concentration increased about 1.9-fold when the incu-
bation time increased from 24 to 48 h at 30 °C and 37 °C.
The quantified proteins and carbohydrates, in biofilm ma-
trices, were not influenced by the growth temperature under
an incubation time of 24 h whatever the surface used (Fig. 2).
However, the increase of the incubation time to 48 h resulted
in a significant increase of matrix production whatever the
temperature used (Fig. 2). By contrast to biofilms grown
during 24 h, increasing growth temperature from 20 °C to
37 °C significantly increased the matrix production for
biofilms grown during 48 h on both surfaces (P<0.05).
Effect of growth temperature, incubation time and surface
type on membrane fatty acids of sessile S. aureus cells
The characterization of membrane fatty acid profiles was
realized for S. aureus grown on stainless steel and polycar-
bonate, at 20 °C, 30 °C and 37 °C, with an incubation time of
Tabl e 4 Total proteins and carbohydrates in the matrix of Staphylococcus aureus biofilms
μg/cm
2
Growth temperature 20 °C 30 °C 37 °C
Times of incubation 24 h 48 h 24 h 48 h 24 h 48 h
Total proteins, SS 11.2± 0.7 18.9± 2.1 10.3±0.8 18.3±1.1 12.1±1.2 24.1±1.9
Total proteins, PC 10.7± 0.6 18.0±1.2 17.9±1.8 19.8±3.5 14.4±1.7 30.0±0.9
Total carbohydrates, SS 4.7± 0.8 5.4± 0.8 4.6± 0.5 8.8± 0.8 6.6± 0.5 12.7±1.5
Total carbohydrates, PC 5.3±1.1 9.1±1.3 5.9±1.3 10.7±1.3 7.6±1.9 12.0± 1.4
Total matrix, SS 15.3± 1.5 24.3± 2.7 14.9±0.6 27.1±11.8 18.7± 0.8 36.8±2.3
Total matrix, PC 16.0±1.5 27.2±1.9 23.8±2.3 30.5±4.5 22.0±1.9 42.0± 1.9
Biofilms were grown on stainless steel (SS) and polycarbonate (PC) substrates, at 20 °C, 30 °C and 37 °C, during 24 and 48 h. The data represent the
mean of the concentration (μg/cm
2
) ± SEM
2602 Appl Microbiol Biotechnol (2014) 98:2597–2607
24 and 48 h (Fig. 3). This investigation was performed to
study the relationship between the membrane fluidity of ses-
sile cells and the biofilm resistance to biocides used in this
work (Fig. 3). Our findings show that the increase of growth
temperature, from 20 °C to 37 °C, resulted in a significant
increase of the total anteiso-branched fatty acids (aBFA) from
56.5 % to 62.8 % (P<0.05), in addition to a decrease of the
total iso-branched fatty acids (iBFA) from 25.3 % to 21.5 %
(P<0.05) for sessile cells grown on stainless steel (Fig. 3). The
changes observed for the aBFA were not related to these of
aC15:0 amounts, which remained at a stable level as shown in
Fig. 3. No significant changes were measured for the aBFA
whatever the temperature used when biofilms were grown on
polycarbonate (Fig. 3). The results of Fig. 3also show that the
aC15:0/aC17:0 ratios were similar whatever the growth tem-
perature, the type of surface and the incubation time used.
However, the increase of the growth temperature from 20 °C
to 37 °C resulted in a significant decrease of the aC17:0/
aC19:0 ratios from 5.9 to 2.8 (P<0.05) in the stainless steel
sessile cells and from 6.9 to 2.6 (P<0.05) in polycarbonate
sessile cells. Moreover, the total saturated fatty acids (SFA)
amount of 24 h sessile cells, remained at a similar level despite
the increase of growth temperature (Fig. 3).
Figure 3also shows that the increase of the incubationtime,
for cells growing under biofilm state from 24 to 48 h, resulted
in a significant increase of aBFAwhatever the temperature and
the surface studied (P<0.05). This could be shown by the
increase in aBFA in stainless steel sessile cells from 56.5 % to
Fig. 2 Total concentration of
proteins and carbohydrates in the
matrix of biofilms grown during
24 h (light gray column) and 48 h
(black column), at 20 °C, 30 °C
and 37 °C, on stainless steel (a)
and polycarbonate (b): data
represent the sum of protein and
carbohydrate concentrations ±
SEM
Fig. 3 Effect of growth temperature, incubation time and surface type on
sessile cells membrane fatty acids profile. a,b: 24 h; c,d:48h.SS
stainless steel, PC polycarbonate, a anteiso,iiso,iBFA total iso-
branched fatty acids, aBFA total anteiso-branched fatty acids, SFA total
saturated fatty acids. Data represent the mean of relative amounts of fatty
acids ± SEM and ratios ± SEM
Appl Microbiol Biotechnol (2014) 98:2597–2607 2603
62.2 % when the incubation time was increased from 24 to
48 h at 20 °C (Fig. 3). The increase of aBFAwas mainly due to
theincreaseofaC15:0(Fig.3). The increase of the growth
temperature from 20 °C to 37 °C, for the biofilm grown for
48 h, showed that the aBFA amount increased from 62.7 % to
70.7 % and from 60.4 % to 68.5 % on the stainless steel and
polycarbonate, respectively (Fig. 3). These increases were the
result of the increase of both aC15:0 and aC19:0 amounts.
These changes resultedin a significantdecrease of the aC17:0/
aC19:0 ratios for sessile cells, from 5.5 to 2.7 on the stainless
steel and from 4.7 to 3.4 on the polycarbonate (P<0.05)
(Fig. 3). In addition, the iBFA and SFA of sessile S. aureus
cells decreased with the increase of the incubation time from
24 to 48 h. This could be shown by the 3 % decrease of the
amount of iBFA on both surfaces (Fig. 3). The decrease of
SFA was ranged between 1.5 and 5.7 % (Fig. 3).
Discussion
The pathogenicity of the food-borne S. aureus is associated
with the ability of this bacterium to produce many toxins in the
food matrix (Normanno et al. 2007). The food contaminations
occur either by food handlers or by the equipment and sur-
faces on which foods are prepared (Simões et al. 2010). In
addition, the food area constitutes a suitable environment for
the biofilm development and several studies have shown the
persistence of S. aureus biofilms on food contact surface
despite the use of cleaning procedures (Gounadaki et al.
2008; Gutierrez et al. 2012). Thus, the purpose of this work
was to set up S. aureus biofilms under different environmental
conditions commonly met during the food processing, such as
temperature change, incubation time and surface type, and to
study their effects on the biofilm resistance to disinfectants.
Moreover, in our study the temperatures used for precultures,
the cultures and the biofilm formation were the same in order
to pre-adapt the cell physiology to these temperatures before
the biofilm formation. However, in many previous studies the
precultures and cultures were carried out under optimal
growth temperature, by contrast to that used for the biofilm
formation (Da Silva Meira et al. 2012;Rodeetal.2007).
Our results show that there is no clear evidence of the effect
of growth temperature on the final biomass of S. aureus
biofilms after 24 and 48 h. These results are in agreement
with what has been reported by Da Silva Meira et al. (2012),
who showed that the growth temperature did not influence the
biofilm formation of S. aureus. However, other studies
underlined that the growth temperature may have an effect
on the biofilm formation of S. aureus with opposite results.
Rode et al. (2007) found that the suboptimal growth temper-
ature (46 °C) increased the biofilm formation of S. aureus.
Choi et al. (2013) and Vazquez-Sanchez et al. (2013)found
that the biomasses of S. aureus biofilms grown at 37 °C were
more important than those of 25 °C. However, Pagedar et al.
(2010) reported a higher cell count of S. aureus biofilms at
25 °C than at 37 °C. Our finding showed also that the stainless
steel and polycarbonate have no significant effect on the
biofilm formation of S. aureus whatever the growth tempera-
ture used. This is in agreement with the results reported by
other studies, which underlined that the surface type did not
affect the biofilm formation of S. aureus (Da Silva Meira et al.
2012; Marques et al. 2007;Songetal.2012). The increase of
the biofilm incubation time did not affect the bacterial biomass
of S. aureus on both surfaces. By contrast, Bae et al. (2012)
showed that the increase of the incubation time resulted in a
decrease of S. aureus biofilm cell density. The discrepancy in
the results, related to the effect of the environmental condi-
tions on the biofilm formation, probably reflects differences in
the experimental conditions, culture medium, surface type and
strains used in each study.
Our results showed that the bacterial biomass was not
influenced by the growth temperature, biofilm age and surface
type, then the comparison of biofilm resistance to disinfectant
products was undertaken. The results of antibiofilm tests
showed that the effect of growth temperature on the biofilm
resistance was related to the incubation time and to the disin-
fectant products used. Increasing growth temperature in-
creased the resistance of the 24-h biofilms to all disinfectants
containing quaternary ammonium chloride (QACs) and
polyhexamethylene biguanides (PHMBs) as active agents.
However, the susceptibility of the 48 h biofilms to P2, con-
taining didecyldimethylammonium chloride (DDAC) as ac-
tive agent, decreased with the increase of growth temperature.
The present findings seem to be consistent with those of
Belessi et al. (2011), who found that the resistance of biofilms
is influenced by the growth temperature. Our findings are also
in agreement with the results of Nguyen and Yuk (2013), who
underlined that the culture conditions such as the temperature,
pH and incubation time may have positive or negative effects
on the resistance of biofilms to disinfectants. Our findings also
showed that the product efficiency depends on the surface
used. The effect of surface could be related to the effect of
surface topography on the biofilm shield and architecture
(Singh et al. 2011), or to its impact on the effectiveness of
cleaning and disinfecting (Chaturongkasumrit et al. 2011;
Schlisselberg and Yaron 2013). The P2 and P3 present 137.5
and 80 ppm of DDAC, respectively. However, the results of
antibiofilm assays showed that the P3 product was more
effective in the killing of sessile cells, grown on the stainless
steel for 24 h, than the P2 products. In addition to DDAC, the
P3 product presents 150 ppm of perfumecontaining morethan
47 % and 7 % of limonene and linalool, respectively. These
compounds are present in many essential oils which have been
found to be effective against pathogenic bacteria such as the
methicillin-resistant S. aureus (Warnke et al. 2009). Thus, the
results may suggest that the essential oil, which are sustainable
2604 Appl Microbiol Biotechnol (2014) 98:2597–2607
and available worldwide, could be a suitable solution to
reduce the dose of chemical disinfectants.
Usually, the differencesin bacterial genera, species, strains,
growth temperatures, disinfectants and incubation times used
in each study makes difficult the comparison between results.
Nevertheless, these studies with our findings highlight the
deep effect of environmental conditions and the bacterial
ecological history on the biofilm resistance. In order to check
the involvement of the biofilm matrix in the S. aureus biofilm
resistance, the total amount of proteins and carbohydrates in
the biofilm matrix were quantified. After 24 h, the
exopolymeric substances production remained unchanged
with the rise of growth temperature. This suggests that biofilm
matrix is probably not involved in the biofilm resistance to
disinfectants. Therefore, our findings highlight that other
mechanisms are maybe involved in the resistance of
S. aureus sessile cells to disinfectant agents. Campanac et al.
(2002), also reported that the resistance of S. aureus biofilms
to QAC could be attributed to phenotypic modifications of
cells rather than the protective presence of an EPS matrix.
Our findings also showed that the increase of the incuba-
tion time to 48 h increased the amount of EPS production in
the matrix of biofilms whatever the growth temperature and
this increase may be involved in the resistance of biofilm to
P2 at 20 °C and to P5 at 30 °C. These results are in agreement
with previous studies which reported that penetration of QAC
in biofilms is hindered and this is probably due to the interac-
tions with EPS of biofilm matrix which lead to an increase of
biofilm resistance (Bridier et al. 2011b; Davison et al. 2010).
The increase of the growth temperature of biofilm, under an
incubation of 48 h, increased the EPS production on both
surfaces. Thus, these results point out again and confirm the
involvement of another factor in the biofilm resistance to
disinfectants, since the biofilms grown during 48 h increased
their sensitivity to P2 disinfectant with the increase of growth
temperature.
It’s well known that the cytoplasmic bacterial membrane is
the first target or barrier to membrane active agents such as the
QACs and PHMB. It has been reported that the hydrophilic
moiety of QACs interacts with the negative charge of the cell
membrane, while the hydrophobic tail strongly interacts with
membrane fatty acids, causing the rearrangement and mem-
brane damage (Gilbert and Moore 2005). The PHMB interact
superficially with the negatively charged acidic phospho-
lipids, inducing its aggregation and therefore disruption of
the lipid bilayer (Gilbert and Moore 2005). Thus, the mem-
brane fatty acids composition, which control the membrane
fluidity, may also play a role in the bacterial resistance to
membrane active agents. Otherwise, several studies have re-
ported the effect of environmental conditions on the modifi-
cations of the membrane fatty acid profiles (Chihib et al. 2003,
2005;Denichetal.2003;Neunlistetal.2005). Our results
underlined the influence of both growth temperature and
incubation time on the membrane fatty acids of sessile
S. aureus cells. The increase of growth temperature of the
biofilm grown during 24 h showed that S. aureusincreased the
amount of aC19:0 which has relatively high melting point and
maintained the aC15:0 and aC17:0 fatty acids at steady state.
The phase transition temperature of the phosphatidylcholine
containing aC19:0 is 36.7 °C which is significantly higher
than those of phosphatidylcholine containing aC17:0 (9.2 °C)
and aC15:0 (−13.9 °C). Thus, the switch to a fatty acid profile
dominated by aC15:0 and aC17:0 and the increase of the
aC19:0 level, in sessile cells when the growth temperature
increased, will probably result in a decrease of the membrane
fluidity. This decrease could be involved in the increase of the
observed resistance of biofilms, grown during 24 h, with the
increase of growth temperature.
When the incubation time increased from 24 to 48 h, our
results showed that the sessile cells increased their membrane
fluidity trough the increase of aBFA (particularly the aC15:0)
and the decrease of SFA relative amount. In fact, the aC15:0
has been reported to be a major determinant of membrane
fluidity for many Gram positive bacteria regarding its low
melting point (Kaneda 1991). In addition, the straight-chain
SFA, are linear and are also known to pack together to produce
a bilayer with a high phase transition (Zhang and Rock 2008).
It has been reported that Gram positive bacteria including
S. aureus maintain high relative amount of aBFA in order to
resist to the environmental stress such as the oxidative one
(Singh et al. 2008). Otherwise, Arce Miranda et al. (2011)
recently showed that the increase of biofilm age of S. aureus
from 24 to 48 h induced oxidative stress in biofilms. Taken
together these two findings, the increase of aBFA observed in
our study is probably due to the reaction of S. aureus to the
increase of oxidative stress in the 48 h biofilms. Increasing
growth temperature from 20 to 37 °C during 48 h increased
also the aBFAs and therefore the membrane fluidity of sessile
S. aureus. These changes in the membrane fluidity may ex-
plain the decrease of the resistance of biofilm grown during
48 h to P2 product when the growth temperature increased.
The increase of biofilm resistance at 20 °C, and the unchanged
resistance of biofilm grown at 30 °C and 37 °C to P2 products,
with the increase of biofilm age suggest that both extracellular
matrix and membrane fluidity may be involved in the resis-
tance of S. aureus biofilm to disinfectant products.
In conclusion, our results showed that the resistance of
biofilm was dependent on several environmental factors com-
monly found in the food sector. It is therefore of interest to be
aware about the ecological history of bacteria and the environ-
mental conditions of the biofilm formation such as tempera-
ture, surface type and biofilm age which may change the
biofilm resistance. Thus, this study suggest that the assessment
of disinfectant efficiency should be carried out against biofilms
formed under different environmental conditions, in order to
decrease the microbiological risk related to their persistence.
Appl Microbiol Biotechnol (2014) 98:2597–2607 2605
Acknowledgments The authors are grateful to the French Agency for
Research and Technology (ANRT) and SCIENTIS laboratory for the
CIFRE grant supporting this work (CIFRE: 2010/0205).
References
Alvarez-Ordonez A, Fernandez A, Lopez M, Arenas R, Bernardo A
(2008) Modifications in membrane fatty acid composition of
Salmonella typhimurium in response to growth conditions and their
effect on heat resistance. Int J Food Microbiol 123(3):212–219. doi:
10.1016/j.ijfoodmicro.2008.01.015
Arad E, Navon-Venezia S, Gur E, Kuzmenko B, Glick R, Frenkiel-
Krispin D, Kramer E, Carmeli Y, Barnea Y (2013) Novel rat model
of methicillin-resistant Staphylococcus aureus-infected silicone
breast implants: a study of biofilm pathogenesis. Plast Reconstr
Surg 131(2):205–214. doi:10.1097/PRS.0b013e3182778590
Arce Miranda JE, Sotomayor CE, Albesa I, Paraje MG (2011) Oxidative
and nitrosative stress in Staphylococcus aureus biofilm. FEMS
Microbiol Lett 315(1):23–29. doi:10.1111/j.1574-6968.2010.02164.x
Bae YM, Baek SY, Lee SY (2012) Resistance of pathogenic bacteria on
the surface of stainless steel depending on attachment form and
efficacy of chemical sanitizers. Int J Food Microbiol 153(3):465–
473. doi:10.1016/j.ijfoodmicro.2011.12.017
Belessi CE, Gounadaki AS, Psomas AN, Skandamis PN (2011)
Efficiency of different sanitation methods on Listeria
monocytogenes biofilms formed under various environmental con-
ditions. Int J Food Microbiol 1(145):25. doi:10.1016/j.ijfoodmicro.
2010.10.020
Bradford MM (1976) A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye
binding. Anal Biochem 72:248–254. doi:10.1016/0003-2697(76)
90527-3
Bridier A, Briandet R, Thomas V, Dubois-Brissonnet F (2011a)
Resistance of bacterial biofilms to disinfectants: a review.
Biofouling 27(9):1017–1032. doi:10.1080/08927014.2011.626899
Bridier A, Dubois-Brissonnet F, Greub G, Thomas V, Briandet R (2011b)
Dynamics of the action of biocides in Pseudomonas aeruginosa
biofilms. Antimicrob Agents Chemother 55(6):2648–2654. doi:10.
1128/AAC.01760-10
Brooks JD, Flint SH (2008) Biofilms in the food industry: problems and
potential solutions. Int J Food Sci Technol 43(12):2163–2176. doi:
10.1111/j.1365-2621.2008.01839.x
Campanac C, Pineau L, Payard A, Baziard-Mouysset G, Roques C
(2002) Interactions between biocide cationic agents and bacterial
biofilms. Antimicrob Agents Chemother 46(5):1469–1474. doi:10.
1128/AAC.46.5.1469-1474.2002
Carpentier B, Cerf O (1993) Biofilms and their consequences, with
particular reference to hygiene in the food industry. J Appl
Bacteriol 75(6):499–511. doi:10.1111/j.1365-2672.1993.tb01587.x
Chaturongkasumrit Y, Takahashi H, Keeratipibul S, Kuda T, Kimura B
(2011) The effect of polyesterurethane belt surface roughness on
Listeria monocytogenes biofilm formation and its cleaning efficien-
cy. Food Control 22(12):1893–1899. doi:10.1016/j.foodcont.2011.
04.032
Chavant P, Gaillard-Martinie B, Hebraud M (2004) Antimicrobial effects
of sanitizers against planktonic and sessile Listeria monocytogenes
cells according to the growth phase. FEMS Microbiol Lett 236(2):
241–248. doi:10.1111/j.1574-6968.2004.tb09653.x
Chihib NE, Ribeiro da Silva M, Delattre G, Laroche M, Federighi M
(2003) Different cellular fatty acid pattern behaviours of two strains
of Listeria monocytogenes Scott A and CNL 895807 under different
temperature and salinity conditions. FEMS Microbiol Lett 218(1):
155–160. doi:10.1111/j.1574-6968.2003.tb11512.x
Chihib NE, Tierny Y, Mary P, Hornez JP (2005) Adaptational changes in
cellular fatty acid branching and unsaturation of Aeromonas species
as a response to growth temperature and salinity. Int J Food
Microbiol 102(1):113–119. doi:10.1016/j.ijfoodmicro.2004.12.005
Choi N-Y, Kim B-R, Bae Y-M, Lee S-Y (2013) Biofilm formation,
attachment, and cell hydrophobicity of foodborne pathogens under
varied environmental conditions. J Korean Soc Appl Biol Chem
56(2):207–220. doi:10.1007/s13765-012-3253-4
Costerton JW, Stewart PS, Greenberg EP (1999) Bacterial biofilms: a
common cause of persistent infections. Science 284(5418):1318–
1322. doi:10.1126/science.284.5418.1318
Crago B, Ferrato C, Drews SJ, Svenson LW, Tyrrell G, Louie M (2012)
Prevalence of Staphylococcus aureus and methicillin-resistant
S. aureus(MRSA) in food samples associated with foodborne illness
in Alberta, Canada from 2007 to 2010. Food Microbiol 32(1):202–
205. doi:10.1016/j.fm.2012.04.012
da Silva Meira QG, de Medeiros Barbosa I, Alves Aguiar Athayde AJ, de
Siqueira-Júnior JP, de Souza EL (2012) Influence of temperature
and surface kind on biofilm formation by Staphylococcus aureus
from food-contact surfaces and sensitivity to sanitizers. Food
Control 25(2):469–475. doi:10.1016/j.foodcont.2011.11.030
Davison WM, Pitts B, Stewart PS (2010) Spatial and temporal patternsof
biocide action against Staphylococcus epidermidis biofilms.
Antimicrob Agents Chemother 54(7):2920–2927. doi:10.1128/
AAC.01734-09
Denich TJ, Beaudette LA, Lee H, Trevors JT (2003) Effect of selected
environmental and physico-chemical factors on bacterial cytoplas-
mic membranes. J Microbiol Methods 52(2):149–182. doi:10.1016/
S0167-7012(02)00155-0
Donlan RM (2002) Biofilms: microbial life on surfaces. Emerg Infect Dis
8(9):881–890
Dubois M, Gilles KA, Hamilton JK, Rebers PA, Smith F (1956)
Colorimetric method for determination of sugars and related sub-
stances. Anal Chem 28(3):350–356. doi:10.1021/ac60111a017
Dubois-Brissonnet F, Naitali M, Mafu AA, Briandet R (2011) Induction
of fatty acid composition modifications and tolerance to biocides in
Salmonella enterica serovar Typhimurium by plant-derived ter-
penes. Appl Environ Microbiol 77(3):906–910. doi:10.1128/AEM.
01480-10
Gilbert P, Moore LE (2005) Cationic antiseptics: diversity of action under
a common epithet. J Appl Microbiol 99(4):703–715. doi:10.1111/j.
1365-2672.2005.02664.x
Gounadaki AS, Skandamis PN, Drosinos EH, Nychas GJ (2008)
Microbial ecology of food contact surfaces and products of small-
scale facilities producing traditional sausages. Food Microbiol
25(2):313–323. doi:10.1016/j.fm.2007.10.001
Gutierrez D, Delgado S, Vazquez-Sanchez D, Martinez B, Cabo ML,
Rodriguez A, Herrera JJ, Garcia P (2012) Incidence of
Staphylococcus aureus and analysis of associated bacterial commu-
nities on food industry surfaces. Appl Environ Microbiol 78(24):
8547–8554. doi:10.1128/AEM.02045-12
Herrera JJ, Cabo ML, Gonzalez A, Pazos I, Pastoriza L (2007) Adhesion
and detachment kinetics of several strains of Staphylococcus aureus
subsp. aureus under three different experimental conditions. Food
Microbiol 24(6):585–591. doi:10.1016/j.fm.2007.01.001
Jenkinson HF, Lappin-Scott HM (2001) Biofilms adhere to stay. Trends
Microbiol 9(1):9–10. doi:10.1016/S0966-842X(00)01891-6
Joseph B, Otta SK, KarunasagarI(2001)Biofilmformationby
Salmonella spp. on food contact surfaces and their sensitivity to
sanitizers. Int J Food Microbiol 64(3):367–372. doi:10.1016/S0168-
1605(00)00466-9
Kaneda T (1991) Iso- and anteiso-fatty acids in bacteria: biosynthesis,
function, and taxonomic significance. Microbiol Rev 55(2):288–
302
Kerouanton A, Hennekinne JA, Letertre C, Petit L, Chesneau O,
Brisabois A, De Buyser ML (2007) Characterization of
2606 Appl Microbiol Biotechnol (2014) 98:2597–2607
Staphylococcus aureus strains associated with food poisoning out-
breaks in France. Int J Food Microbiol 115(3):369–375. doi:10.
1016/j.ijfoodmicro.2006.10.050
Marques SC, Rezende JGOS, Alves LAF, Silva BC, Alves E, Abreu LR,
Piccoli RH (2007) Formation of biofilms by Staphylococcus aureus
on stainless steel and glass surfaces and its resistance to some
selected chemical sanitizers. Braz J Microbiol 38:538–543. doi:10.
1590/S1517-83822007000300029
Neunlist MR, Federighi M, Laroche M, Sohier D, Delattre G, Jacquet C,
Chihib NE (2005) Cellular lipid fatty acid pattern heterogeneity
between reference and recent food isolates of Listeria
monocytogenes as a response to cold stress. Antonie Van
Leeuwenhoek 88(3–4):199–206. doi:10.1007/s10482-005-5412-7
Nguyen HDN, Yuk H-G (2013) Changes in resistance of Salmonella
Typhimurium biofilms formed under various conditions to industrial
sanitizers. Food Control 29(1):236–240. doi:10.1016/j.foodcont.
2012.06.006
Normanno G, La Salandra G, Dambrosio A, Quaglia NC, Corrente M,
Parisi A, Santagada G, Firinu A, Crisetti E, Celano GV (2007)
Occurrence, characterization and antimicrobial resistance of entero-
toxigenic Staphylococcus aureus isolated from meat and dairy prod-
ucts. Int J Food Microbiol 115(3):290–296. doi:10.1016/j.
ijfoodmicro.2006.10.049
Pagedar A, Singh J, BatishVK (2010) Surface hydrophobicity, nutritional
contents affect Staphylococcus aureus biofilms and temperature
influences its survival in preformed biofilms. J Basic Microbiol
50(1):201000034. doi:10.1002/jobm.201000034
Poulsen LV (1999) Microbial Biofilm in food processing. LWT-Food Sci
Technol 32(6):321–326. doi:10.1006/fstl.1999.0561
Rode TM, Langsrud S, Holck A, Moretro T (2007) Different patterns of
biofilm formation in Staphylococcus aureus under food-related
stress conditions. Int J Food Microbiol 116(3):372–383. doi:10.
1016/j.ijfoodmicro.2007.02.017
Schlisselberg DB, Yaron S (2013) The effects of stainless steel finish on
Salmonella Typhimurium attachment, biofilm formation and sensi-
tivity to chlorine. Food Microbiol 35(1):65–72. doi:10.1016/j.fm.
2013.02.005
Simões M, Simões LC, Vieira MJ (2010) A review of current and
emergent biofilm control strategies. LWT-Food Sci Technol 43(4):
573–583. doi:10.1016/j.lwt.2009.12.008
Singh VK, Hattangady DS, Giotis ES, Singh AK, Chamberlain NR,
Stuart MK, Wilkinson BJ (2008) Insertional inactivation of
branched-chain alpha-keto acid dehydrogenase in Staphylococcus
aureus leads to decreased branched-chain membrane fatty acid
content and increased susceptibility to certain stresses. Appl
Environ Microbiol 74(19):5882–5890. doi:10.1128/AEM.00882-08
Singh AV, Vyas V, Patil R, Sharma V, Scopelliti PE, Bongiorno G,
Podesta A, Lenardi C, Gade WN, Milani P (2011) Quantitative
characterization of the influence of the nanoscale morphology of
nanostructured surfaces on bacterial adhesion and biofilm forma-
tion. PLoS One 6(9):26. doi:10.1371/journal.pone.0025029
Song L, Wu J, Xi C (2012) Biofilms on environmental surfaces: evalu-
ation of the disinfection efficacy of a novel steam vapor system. Am
J Infect Control 40(10):926–930. doi:10.1016/j.ajic.2011.11.013
Srinivasan A, Wolfenden LL, Song X, Mackie K, Hartsell TL, Jones HD,
Diette GB, Orens JB, Yung RC, Ross TL, Merz W, Scheel PJ,
Haponik EF, Perl TM (2003) An outbreak of Pseudomonas
aeruginosa infections associated with flexible bronchoscopes. N
Engl J Med 348(3):221–227. doi:10.1056/NEJMoa021808
To MS, Favrin S, Romanova N, Griffiths MW (2002) Postadaptational
resistance to benzalkonium chloride and subsequent physicochem-
ical modifications of Listeria monocytogenes. Appl Environ
Microbiol 68(11):5258–5264. doi:10.1128/AEM.68.11.5258-5264.
2002
Tote K, Horemans T, Vanden Berghe D, Maes L, Cos P (2010) Inhibitory
effect of biocides on the viable masses and matrices of
Staphylococcus aureus and Pseudomonas aeruginosa biofilms. Appl
Environ Microbiol 76(10):3135–3142. doi:10.1128/AEM.02095-09
Vazquez-Sanchez D, Habimana O, Holck A (2013) Impact of food-
related environmental factors on the adherence and biofilm forma-
tion of naturalStaphylococcus aureusisolates. Curr Microbiol 66(2):
110–121. doi:10.1007/s00284-012-0247-8
Warnke PH, Becker ST, Podschun R, Sivananthan S, Springer IN, Russo
PA, Wiltfang J, Fickenscher H, Sherry E (2009) The battle against
multi-resistant strains: renaissance of antimicrobial essential oils as a
promising force to fight hospital-acquired infections. J
Craniomaxillofac Surg 37(7):392–397. doi:10.1016/j.jcms.2009.
03.017
Zhang YM, Rock CO (2008) Membrane lipid homeostasis in bacteria.
Nat Rev Microbiol 6(3):222–233. doi:10.1038/nrmicro1839
Appl Microbiol Biotechnol (2014) 98:2597–2607 2607
