Content uploaded by Thomas Eugene Cloete
Author content
All content in this area was uploaded by Thomas Eugene Cloete on Jul 18, 2014
Content may be subject to copyright.
African Journal of Microbiology Research Vol. 4(14), pp. 1515-1524, 18 July, 2010
Available online http://www.academicjournals.org/ajmr
ISSN 1996-0808 ©2010 Academic Journals
Full Length Research Paper
Protease and amylase enzymes for biofilm removal and
degradation of extracellular polymeric substances
(EPS) produced by Pseudomonas fluorescens bacteria
I. Phyllis Molobela1*, T. Eugene Cloete2 and Mervyn Beukes3
1Department of Microbiology and Plant Pathology, University of Pretoria, Pretoria, South Africa.
2Faculty of Science, University of Stellenbosch, South Africa.
3Department of Biochemistry, University of Pretoria, Pretoria, South Africa.
Accepted 8 June, 2010
Removal of biofilms is difficult. In industrial settings, both the inactivation and removal of biofilms are
of huge concern. If only disinfection without the removal of attached biofilms occurs, the inactivated
biofilm cells may provide an ideal environment for further adhesion and growth, resulting in a complex
matrix. Microbial resistance to biocides and their negative environmental impact are the main reasons
for finding alternative biofilm control strategies. Enzymes may offer such an alternative. The objective
of this study was to determine the effect of commercial proteases and amylases on biofilms formed by
Pseudomonas fluorescens. Biofilms were grown in diluted medium containing glass wool used as the
attachment surface. Extracellular polymeric substances (EPS) were extracted and EPS composition was
determined. Protease (savinase, everlase and polarzyme) and amylase (Amyloglucosidase and Bacterial
Amylase Novo) activity was tested on both biofilms and on extracted EPS. After testing enzymes,
biofilm integrity was evaluated by scanning electron microscopy. EPS composition consisted
predominantly of proteins. Everlase and Savinase were the most effective enzymatic treatments on
removing biofilms and degrading the EPS.
Key words: Biofilms, extracellular polymeric substances, Pseudomonas fluorescens, proteases, amylases.
INTRODUCTION
When bacterial cells approach inert surfaces, they first
bound to the substratum by weak forces involving their
external structures such as flagella, fimbriae or capsular
components (Xavier et al., 2005). As the cells remain
attached to the surface for some time, they secret sticky
extracellular polymeric substances (EPS) forming a
biofilm matrix that embeds several layers of bacterial
cells once the biofilms mature (Orgaz et al., 2006;
Flemming et al., 2007). EPS are composed of a wide
variety of materials including polysaccharides, proteins
(Johansen et al., 1997; Leroy et al., 2008), nucleic acid,
uronic acid and humic substances (Orgaz et al., 2006).
Polysaccharides are partly responsible for bacterial
adhesion and biofilm accumulation on the surface
*Corresponding author. E-mail: mantlophyllis@yahoo.com. Tel:
+27 12 420 2995. Fax: +27 12 420 3266.
(Loiselle et al., 2003). The EPS also serves many other
functions such as providing an adhesive foundation,
structural integrity, bacterial protection and intercellular
communication (Zhang et al., 2005; de Carvalho, 2007;
Ploux et al., 2007; Leroy et al., 2008).
The difference in the quantity of biofilm EPS is a result
of the growing conditions of the biofilms (O’ Toole et al.,
2000). EPS has a complex architectural structure
(Flemming et al., 1998) containing channels which allow
the inflow of water, oxygen and nutrients and outflow of
byproducts (Zhang et al., 2001; Arevalo-Ferro et al.,
2005; Donlan, 2002) and enhances bacterial resistance
to antimicrobial agents(Parkar et al., 2000; Prakash et al.,
2003; Lequette et al., 2010).
EPS serves other functions including: Facilitation of the
initial attachment of bacterial cells to a surface (Stoodley
et al., 2002); Formation and maintenance of the micro
colony (Flemming et al., 1998); Enables the bacteria to
capture nutrients (Gomez-Suarez et al., 2002) causes
1516 Afr. J. Microbiol. Res.
biofouling (Cloete et al., 1998); Facilitates cell-cell
communication (Zhang et al., 2001) and also function as
a stabilizer of the biofilm structure and as a barrier
against hostile environments (Zhang et al., 2001;
Arevalo-Ferro et al., 2005; Lapidot et al., 2006; Ploux et
al., 2007; Donlan, 2002).
The production of EPS is influenced by internal and
external factors including: Quorum sensing (cell to cell
communication); Surface topography, hydrodynamic
shear forces; Fluid velocity and nutrient availability
(Cloete, 1998; Cloete, 2003; Sreenivasan et al., 2005).
EPS is a complex structure made up of different
components including carbohydrates, proteins, lipids and
nucleic acid (Flemming, 1998, Allison et al., 2000; Liu et
al., 2003).
Previous studies have indicated that disinfection with
chlorine dioxide and chlorine, for example, can reduce
the concentrations of planktonic bacteria, but have little to
no effect on the concentrations of biofilm bacteria (Berry
et al., 2006). The mechanism behind the resistance of
biofilms to disinfection is through protection of the biofilm
cells that are embedded in the extracellular polymeric
substances (Xavier et al., 2005; Walker et al., 2007).
Enzymes have been used and proven to be effective
for the degradation of the multistructural EPS of the
biofilms (Johansen et al., 1997; Melo et al., 1997;
Augustin et al., 2004; Lequette et al., 2010). The mode in
which enzymes destroy the EPS is by degrading the
physical integrity of the EPS (Xavier et al., 2005). Walker
et al. (2007) indicated that in order to design enzymes
that target the EPS of the biofilms, it is important to have
an understanding of the nature of the EPS. The efficiency
of any one enzyme degrading EPS will depend on the
EPS composition (Xavier et al., 2005; Walker et al.,
2007).
Previous studies have been published regarding
enzyme degradation of mature biofilms using synthetic
polysaccharides (Loiselle et al., 2003; Vickery et al.,
2004). Cellulase from Penicillium funiculusum was
effective in degrading mature biofilms of Pseudomonas
aeruginosa; and it was also found to be useful in
degrading the exopolysaccharides of Pseudomonas
fluorescens (Loiselle et al., 2003; Vickery et al., 2004).
Therefore, the application of enzymes to degrade EPS is
a promising and an attractive option in many industries
where complete biofilm removal is essential.
The aim of this study was to test selected commercial
proteases and amylases for their effectiveness in the
degradation and removal of EPS produced within a
Pseudomonas fluorescens biofilm. We further establish
some standard protocols for the evaluation of enzyme
efficiency in degrading EPS and biofilm.
MATERIALS AND METHODS
Pseudomonas fluorescens inoculums used for biofilm growth
P. fluorescens was inoculated into sterile Nutrient Broth and
incubated aerobically at 26°C overnight. During the incubation
period, one set of P. fluorescens samples was daily fed with 2 ml of
the diluted medium and the control P. fluorescens samples were
unfed. Bacterial growth was monitored daily by measuring the
optical density at 620 nm. After incubation, the concentration of the
bacterial suspension was adjusted to OD, 620 nm.
Biofilm formation and growth
P. fluorescens was grown according to Rochex and Lebeault (2007)
with modification. A tandardized P. fluorescens suspension (100 µl)
was inoculated into flasks containing 100 ml sterile Nutrient Broth
(Merck) and 2 g of glass wool used as a surface for the growth of
biofilm. Flasks containing the glass wool were incubated at 26°C for
7 days with continuous agitation at 100 rpm. During biofilm growth,
flasks marked CF100XNB were daily fed with 2 ml of Nutrient Broth
and those marked WAN were unfed. At day 7, bacterial suspen-
sions containing the planktonic cells were discarded from the flasks.
Quantitative determination of viable cells
Ten fold series of dilutions were made by inoculating 100 µl of the
bacterial suspensions to 900 µl of Ringer’s solutions and mix. The
aliquots (0.1 ml) were spread onto sterile Nutrient agar plates
(Merck) and incubated for 24 – 48 h at 26°C (3 plates for each
dilution). Viable cells were enumerated and expressed as colony
forming units (CFU/ml).
Microtiter assay for efficacy of biofilm removal
The Microtiter assay was performed according to Pitts et al. (2003)
with the following modifications; 200 µl of standardized bacterial
suspension was added to the wells of a polystyrene microtiter plate
(Lasec, S.A.) and incubated at 26°C with shaking at 100rpm for 48
h. Biofilm formation was monitored periodically by visual inspection.
After incubation, the supernatant was discarded and plates were
washed three times with 200 µl sterile distilled water to remove non
adherent bacterial cells. To each well, 1 U/ml and 2 U/ml of
proteases and amylases were added. A well without enzymes was
used as control. Plates were incubated for 1 h at 26°C. Following
incubation, plates were emptied and washed twice with sterile
distilled water. The remaining cells were fixed with 200 µl of 95%
ethanol for 15 min and allowed to dry. Crystal violet solution (200
µl) was added into each well for 30 min. Plates were washed five
times with sterile distilled water. Wells were washed with 30%
glacial acetic acid (200 µl) (Merck, S.A.). Plates were read at 595
nm using a Multiskan Ascent ELISA plate reader (Termo
Labsystems). The experiment was performed in duplicate.
The micro titer screening method was used to quantitatively
measure the removal efficacy of proteases and amylases on
biofilms of P. fluorescens. A measure of efficacy called Percentage
Reduction by Pitts et al. (2003) was used to evaluate the efficacy of
these enzymes.
Percentage reduction = [(C –B) – (T – B))/ (C – B)] × 100%
Where:
B denotes, the average absorbance per well for blank (no biofilm,
no treatment); C denotes the average absorbance per well for
control wells (biofilm, no treatment) and T denotes the average
absorbance per well for treated wells (biofilm and treatment).
Molobela et al. 1517
Table 1. Properties of the commercial enzymes tested in this study.
Optima conditions
Name Enzyme Manufacturer Source pH Temperature (°C) Application
Savinase Protease Novozyme Genetically modified Bacillus clausi 8 - 11 15 - 75 Laundry
Everlase Protease Novozyme Genetically modified Bacillus clausii 8 - 11 15 - 75 Detergent industry
Polarzyme Protease Novozyme Genetically modified Bacillus spp 9 - 11 20 - 40 Detergent industry
*BAN Alphamylase Novozyme Bacillus amyloliquefaciens 6 - 7 20 - 60 Food industry
¤AMG Glucoamylase Novozyme Aspergillus niger 4 - 5 20 - 60 Food industry
*Bacterial Amylase Novo, ¤Amyloglucosidae.
Biofilm detachment and extraction of extracellular polymeric
substances (EPS)
Flasks containing glass wool with attached biofilm cells were
vortexed vigorously for 5 min to detach loosely bound biofilm cells.
Bacterial aliquot (20 ml) was added to 50 ml sterile centrifuge tubes
(Merck). The contents were homogenized for about 30 s using a
Cole-Parmer homogenizer at an adjusted output of 50% and spun
at 3500 xg for 5 min at 4°C. The supernatants were transferred to
sterile centrifuge tubes and further spun at 9000 xg for 30 min, 4°C.
Pellets were resuspended in 20 ml sterile distilled water, freeze
dried and dissolved in Phosphate and Tris-Maleate buffers, res-
pectively for enzyme assays. Dissolved pellets were filtered through
0.8/0.2 µm filters (Acrodics PF, PA//Inc). Filtrates were assayed for
EPS composition and enzyme activity.
Determination of the carbohydrate concentration in the EPS
The carbohydrate concentration was determined according to
Gaudy’s method (1962). Briefly, pellets were dissolved in
Phosphate and Tris-Maleate buffers (1 ml). Freshly prepared
Anthrone solution (1 ml) was added in each test tube. The mixture
was incubated in a water bath at 95°C for 15 min. After incubation,
the mixture was allowed to cool to room temperature. Cooled
aliquots (200 µl) were transferred to micro plate wells (Lasec, S.A.)
and read at 620 nm using a plate reader (Multiskan Ascent V1.24,
Amersham). Glucose was used as a standard to construct a
standard curve.
Determination of the protein concentration in the EPS
Protein concentration was determined by the modified method of
Lowry (Frøelund et al., 1995). Extracellular polymeric substances
(EPS) (10 µl) were added into wells of a micro titter plate. Control
wells were added with phosphate buffer. Coomassie plus reagent
(300 µl) was added to each well. The plate was incubated at room
temperature for 10 min. After incubation, absorbances were read at
595 nm using a Multiskan Ascent V1.24 plate reader, (Amersham).
Bovine serum albumin (BSA) was used as a standard to construct a
standard curve.
Enzymes used in the study
Activity of the enzymes, listed in Table 1, on biofilm removal was
evaluated for biofilm removal. The proteases were: (1) Savinase,
(2) Everlase and (3) Porlazyme. The Amylases were: (4)
Amyloglucosidase (AMG) and (5) Bacterial Amylo Novo (BAN). All
enzymes used were purchased from Novozymes (Ltd) South Africa.
Proteases were diluted in 0.1 M Phosphate buffer, pH 8.3. Bacterial
Amylo Novo (BAN) was dissolved in 0.2 M Tris-maleate, pH 7.0;
and Amyloglucosidase (AMG) was diluted in Phosphate buffer, pH
5.
ENZYMATIC TREATMENTS
Degradation of biofilm EPS
Following protein and carbohydrate analysis, 1 ml of suspended
EPS was added into 50 ml centrifuge tubes containing the protease
or amylase enzymes diluted in specific buffer solutions. The
samples were incubated at 26°C and aliquots were taken at 15 min
intervals. For the protease activity, 300 µl of sample was transferred
to micro plates and analyzed via the Bradford assay, while the
amylase activity was analyzed using the Anthrone assay.
Testing of enzymes for the removal of biofilm cells on the
glass wool
Glass wool – attached biofilms from fed and unfed cultures were
incubated with enzyme solutions (100 ml) at 26°C for 24 h without
agitation. Biofilms with no enzymes were used as control. After the
incubation period, the effect of enzymatic activity on the biofilms
was evaluated using the Scanning Electron Microscope (SEM).
Sample preparation for Scanning Electron Microscopy
Glass wool samples were fixed in a solution of 2.5% glutaraldehyde
in 75 mM Phosphate buffer, pH 7.4 for 1 h. Samples were rinsed
three times for 15 min at a time in 50% 75 mM phosphate buffer.
After the rinsing step, samples were dehydrated in ethanol at
concentrations of 50, 70, 90 and three times 100% each for 15 min
respectively. After the drying step samples were critically dried with
CO2 (Martin et al., 2006). Samples were coated with gold and
visualized using a Scanning Electron Microscope (JSM-840, JEOL,
TOKYO Japan).
RESULTS
Growth and viable cells of Pseudomonas fluorescens
The rate of P. fluorescens growth was maximal after the
6th day of incubation and progressively reached a plateau
phase thereafter. P. fluorescens growing in the daily fed
medium (CF100XNB) was slightly higher than unfed
1518 Afr. J. Microbiol. Res.
Figure 1. Growth of Pseudomonas fluorescens in fed and unfed nutrient
medium conditions. Bars indicate standard errors.
Table 2. Comparison of viable cells between fed and unfed Pseudomonas fluorescens biofilms.
Viable cells (CFU/ml) ×105 Average ± SD *
Fed P. fluorescens 1.93 ± 8.485
Unfed P. fluorescens 1.76 ± 5.657
*Average ± Standard deviation.
0
10
20
30
40
50
60
70
80
90
100
Percentage Reduction (%)
EVERLASE
SAVINASE
POLARZYME
BAN
AMG
RINGERS
Figure 2. Microtiter assay for the evaluation of enzyme efficacy
for the removal of Pseudomonas fluorescens biofilms. Bars
indicate standard errors.
P. fluorescens (WAN) (Figure 1). Plate count assay
results showed slightly more viable cells in the fed P.
fluorescens growth than the unfed growth of P.
fluorescens (Table 2).
Microtiter assay for the evaluation of enzyme efficacy
for biofilm removal
The micro plate assay was used to determine the activity
of enzymes on the reduction of P. fluorescens biofilms
and on the degradation of extracellular polymeric sub-
stances (EPS). Savinase and Everlase showed a highest
percentage reduction (Figure 2). This was followed by
Amyloglucosidase (AMG) with higher percentage
reduction and Bacterial Amylo Novo was less effective
with lower percentage reduction. Polarzyme was not
effective for removal of P. fluorescens biofilms and was
comparable to the control (biofilms treated with Ringer’s
solution (Figure 2).
EPS, proteins and carbohydrate concentrations
The extracellular polymeric substances (EPS)
concentration of the biofilm that was fed daily (0.219
g/ml) was slightly higher than the EPS concentration in
the unfed biofilms (0.126 g/ml) (Table 3). The protein
concentration in both fed and control experiments were
higher than carbohydrate concentrations in both experi-
ments (Table 3). However, the protein concentration in
Molobela et al. 1519
Table 3. Comparison of the production of extracellular polymeric substances (EPS), protein and carbohydrate
concentrations of the fed and unfed Pseudomonas fluorescens biofilms
Fed biofilms (Av ± SD)* Unfed biofilms (Av ± SD)*
EPS mass
(g/OD 620)
Protein
(µg/ml)
carbohydrate
(µg/ml)
EPS mass
(g/OD 620)
Proteins
(µg/ml)
Carbohydrate
(µg/ml)
0.219 (± 0.001) 1592 (± 1.989) 119.8 (± 0.004) 0.126 (± 0.023) 1474 (± 1.767) 92.2 (± 0.002)
*Average ± Standard deviation.
Figure 3. Degradation potential of a. Savinase b. Everlase c. Polarzyme on extra cellular
polymeric substances (EPS) produced by Pseudomonas fluorescens biofilms. Bars indicate
standard errors.
the EPS of the biofilm cultured with daily feeding (1592
µg/l) was higher than the protein concentration in the
control EPS (1474 µg/l) (Table 3). Similarly, carbohydrate
concentrations were higher in the daily fed (119.8 µg/l)
than control EPS (92.2 µg/l) (Table 3).
Enzymatic degradation of proteins and
carbohydrates in the EPS
The proteases, Savinase and Everlase were the most
effective enzymes for the degradation of protein
concentration of the extracted EPS (Figures 3A and B).
Polarzyme did not show any reduction in the protein
concentration (Figure 3C). The amylase
Amiloglucosidase (AMG) was partially effective while
the amylase Bacterial Amylase Novo (BAN) was the least
effective on carbohydrate degradation in the EPS (Figure
4A and B). The control EPS protein and carbohydrate
concentrations remained unaffected (Figure 4C). Micro-
scopic studies of the effect of enzymes on the P.
fluorescens biofilms revealed that biofilms treated with
Savinase and Everlase showed a reduction in biofilm
cells and substantial degradation of the extracellular
1520 Afr. J. Microbiol. Res.
Figure 4. Degradation potential of: A. Amyloglucosidase B. Bacterial Amylase Novo on extracellular polymeric
substances (EPS) produced by Pseudomonas fluorescens biofilms C. Non treated EPS. Bars indicate standard errors.
polymeric substances (EPS) (Figures 5A and B). The
amylase Amyloglucosidase and Bacterial Amylase Novo
treated biofilms were partially degraded (Figures 5C and
D).
DISCUSSION
The effect of nutrient concentration on biofilm yield
There was a slight difference in the number of viable cells
grown in the fed and unfed nutrient medium conditions
and there was no noticeable difference in biofilms cells
grown in fed and unfed medium but there was a
difference in the amount of EPS produced. The fed
biofilms had more EPS than the unfed biofilms. Nutrients
boosted the biofilm cells growing in rich medium which
resulted in more EPS produced. It was indicated in
previous studies that biofilms growing in high nutrient
medium were more abundant, densely packed and
thicker (Allison et al., 2000; Prakash et al., 2003; Rochex
and Lebeault, 2007).
Rochex and Lebeault (2007) showed that nutrient
conditions influenced biofilm formation of bacterial strains
isolated from a paper machine. Rochex and Lebeault
(2007) also compared biofilms growing in two different
medium concentrations and found that the biofilm mass
in medium containing 0.1 g/l of glucose was 90% lower
than the biofilm mass in medium containing 0.5 g/l of
glucose.
Protein and carbohydrate concentrations in the
biofilm EPS
The EPS of P. fluorescens biofilm grown in fed medium
had a higher protein and carbohydrate concentration than
in the unfed biofilm EPS. Protein concentrations were
higher than the carbohydrate concentration in both fed
and unfed biofilms. This indicated that the structural
components of the biofilm EPS was dependent on the
nutrient status in which the biofilm was grown. These
Molobela et al. 1521
Figure 5. Microscopic analysis of degradation actions of enzymes on extra cellular polymeric
substances (EPS) of Pseudomonas fluorecens biofilms attached on the glass wool fibers after 24
h incubation at 26˚C. A. Savinase B. Everlase, C. Amyloglucosidase D. Bacterial Amylase Novo
E. Polarzyme F. Non treated biofilms.
Results correspond to the work of Simoes (2003) who
found more protein (total protein = 217.7 mg/g) than
carbohydrate (total carbohydrate = 63.3 mg/g) in the EPS
produced by P. fluorescens biofilms under specific growth
conditions.
In some studies, it was indicated that carbohydrates
are the main constituents of the EPS while some studies
found proteins to dominate (Zhang et al., 2001; Liu et al.,
2003; Orgaz et al., 2006). In this study proteins were
found to be dominant rather than carbohydrates. None-
theless, the EPS components of the biofilms differ in
quantity; structure or nature depending on the micro-
organisms within the biofilm.
The structural components of the EPS depend on the
type of microorganisms within the biofilm. Allison et al.
(2000) indicated that the EPS of the biofilms is highly
heterogenous even among the same bacterial species
and therefore its composition and function within the
biofilms will differ. O’ Toole et al. (2000) indicated that
different biofilms produce different amounts of EPS.
In addition, depending on the extraction protocols used,
the EPS composition will differ (Liu et al., 2002; Augustus
and Ali-Vehmas, 2004). Liu et al. (2002) studied mixed
cultures in wastewater treatment systems and found that
the protein (41.3%) concentration was greater than the
carbohydrate concentration (18.7%) in the methanogenic
sludge when the formaldehyde–NaOH extraction method
was applied. In addition, the formaldehyde–NaOH pro-
cess extracted the highest concentration of EPS from all
the sludges. In this study, EPS of the biofilms was ex-
tracted by centrifuging the sample at low and high speed
to separate the biomass from the EPS. This method was
chosen because of its higher extraction efficiency and
lower cell lyses. Then, the Anthrone and Lowry assays
1522 Afr. J. Microbiol. Res.
were employed for the quantification of glucose and
protein concentrations respectively in the EPS. Anthrone
and Lowry assays were employed in this study for the
quantification of total carbohydrate and proteins in the
EPS since enzymes were tested for the degradation of a
broad spectrum of carbohydrates and proteins. These
assays are based on the colorimetric determination of
colour development. The advantage is that these assays
can also be performed in a micro plate format and can be
performed at room temperature. In addition standard
curves can be constructed to convert the absorbencies
into con-centrations.
The use of protease and amylase enzymes for the
degradation of EPS
Many antimicrobial agents fail to penetrate the biofilm
due to the EPS which acts as a barrier protecting the
bacterial cells within. The alternative will be the use of
compounds which can degrade the EPS of the biofilm
(Loiselle et al., 2003; Walker et al., 2007). Enzymes have
been proven to be effective for the degradation of the
EPS of the biofilms (Johansen et al., 1997; Melo et al.,
1997; Lequette et al., 2010). Enzymes remove biofilms
directly by destroying the physical integrity of the EPS
(Liu et al., 2004; Xavier et al., 2005). The mechanism in
which enzymes destroy the physical integrity of the EPS
is through weakening the proteins, carbohydrate and lipid
making up the structures of the EPS through the
degradation process. For efficient removal of biofilm, it is
therefore important that the structural components of the
EPS should be known before application of the relevant
enzymes.
In the present study, enzymes were tested for the
eradication of P. fluorescens biofilms. All enzymes tested
except for the protease Polarzyme were effective for the
degradation of the biofilm EPS. Savinase and Everlase
were the most effective for the degradation of P.
fluorescens EPS. The reason for the inefficiency of
Polarzyme may be due to its incompatibility with the
specific protein structural components of the biofilm EPS
tested in this study. The manner in which the enzymes
degrade the proteins in the EPS is through binding and
hydrolysis of the protein molecules and converting them
into smaller units that can be transported through the cell
membranes and then be metabolized. The mode of enzy-
matic action will therefore depend on the specific protein
structure and this in turn will determine its efficacy.
The multi structural components of the EPS may be
derived from proteins, glycoproteins, nucleic acid,
glycolipid, phospholipids including humic substances
which are non cellular substances (Liu et al., 2004). The
efficiency of the proteases may therefore be due to their
broad spectrum activity in degrading a variety of proteins
acting partly as the multi structural components of P.
fluorescens and mixed bacterial species biofilm EPS.
Extracellularly secreted proteins are substances with
molecular weight between 10 and 200 kDa. These
compounds contain 40 – 60% of hydrophobic amino
acids. It was observed that the extra cellular proteins syn-
thesized by Sulfolobus acidocalcidarius are composed
mostly of amino acid with hydroxyl group. However, the
Bacillus subtilis extracellular protein layer is a com-
position of L and D glutaminosyl residues (Czaczyk and
Myszka, 2007). According to Ton–That et al. (2004) the
ratio of glutaminosyl isomers in Bacillus subtilis extra-
cellular protein layer changed significantly in oxygen
limited conditions.
Leroy et al. (2007) also found the protease, Savinase to
be more effective for the prevention of adhesion and
detachment of a Pseudoalteromonas sp. D14 biofilm than
xylanase, amylase, cellulase and lipase. Ledder et al.
(2008) also found protease to be effective for the removal
of A. naeslundii and F. nucleatum biofilm.
Donlan (2002) indicated that EPS may be hydrophilic or
hydrophobic depending on the structural components
making up such EPS and the environmental conditions
were the biofilms are developing. Studies have indicated
that among one bacterial species EPS components may
differ (Czaczyk et al., 2007). The structure of
polysaccharides synthesized by microbial cells may vary.
Microbial exopolysaccharides are comprised of either
homopolysachharides or heteoropolysaccharides. Hom-
opolysaccharides are composed of only one mono-
saccharide type such as D – glucose or L- fructose
(Czaczyk et al., 2007). Homopolysaccharides belong to
three distinct groups including: – D – glucan which is
produced by Leuconostoc mesenteroides; ß- D- glucans
which is produced by Pediococcus spp. and
Streptococcus spp.; Fructans are produced by
Streptococcus salivarius.
A number of lactic acid bacteria produce heteropoly-
saccharides. These molecules form from repeating units
of monosaccharides including D- glucose, D- galactose,
L- fructose, L- rhamnose, D- glucuronic acid, L- guluronic
acid and D- mannuronic acid. The type of both linkages
between monosaccharides units and the branching of the
chain determines the physical properties of the microbial
heteropolysaccharides (Sutherland, 2001; Czaczyk et al.,
2007). As an example, bacterial alginate is a heteropoly-
saccharide with an irregular structure. In this polymer, D-
mannurosyl and L- guluronosyl residues are found.
Alginate is mostly produced by the cells of Pseudomonas
aeruginosa and Azatobacter vinelandii (Czaczyk et al.,
2007). Due to a wide range of linkages and the
complexity of polysaccharides structures, it would
therefore be difficult for most amylase enzymes (including
the test amylases) to break down the bond linkages of
the monomers making up polysaccharides which
determine the physical structure of the EPS.
It was therefore not surprising that the amylase
enzymes tested for the degradation of P. fluorescens
biofilms, were less effective than the proteases. This is
also in agreement with previous studies, indicating that
the activity of most amylase enzymes tested was less
effective for the removal of bacterial biofilms than
proteases (Ledder et al., 2008). This was attributed to the
dominance of proteins in the EPS. In most cases proteins
seem to be the main constituents of the biofilms EPS and
are found mostly at the outer layer of the biofilms (Liu et
al., 2004; Bhaskar and Bhosle, 2005). Therefore, it is
unlikely that the amylase enzymes would degrade the
protein in the EPS. Since the biofilm EPS was made up
of mostly proteins it explains why the amylase enzymes
were less efficient for biofilm degradation.
Scanning electron microscopy analysis of enzyme
efficiency for EPS degradation
SEM analysis confirmed that protease enzymes
(Savinase and Everlase) were more effective than the
amylase enzymes for degrading the EPS of P.
fluorescens (Figure 4).
Conclusion
If a compound or compounds capable of destroying all
the structural components of different EPS that are
produced by different biofilms growing under different
conditions is found then the “city of microbes” (biofilms)
would be destroyed permanently. If only an enzyme or
enzymatic mixture capable of shutting down or deactivating
the quorum sensing systems of different biofilm EPS
could be found, then there would not be any formation of
biofilms and the name biofilm will undergo extinction.
Enzymes differed in activity. Protease enzymes were
capable of destroying the “house of the microbes” (EPS).
The amylase enzymes were less effective for the
degradation of P. fluorescens biofilms. This may be due
to the fact that EPS is highly heterogeneous even among
the bacteria of the same species and therefore its
structural composition will differ. Another reason for the
difference in enzyme activity may be the way they were
formulated and the mode of action. In conclusion, in order
to design enzymes which target the EPS of the biofilms, it
is important to have an understanding of the structural
composition of the EPS.
ACKNOWLEDGEMENTS
The authors would like to thank, the Laboratory for
Microscopy and Microanalysis (University of Pretoria) for
assisting in sample analysis; Department of Biochemistry,
(University of Pretoria) for technical assistance; financial
support from the University of Pretoria and ESKOM and
National Research Foundation (NRF). Finally, the authors
would like to thank Novozymes South Africa for supplying
the enzymes.
Molobela et al. 1523
REFERENCES
Allison DG (2000). Microbial biofilms: Problems of control. In: Allison
DG, Gilbert P, Lappin – Scott H Wilson M (Eds) Community structure
and cooperation in biofilms.. Cambridge University Press. pp, 309-
327
Arevalo-Ferro C, Reil G, Gorg A, Eberl L, Riedel K (2005). Biofilm
formation of Pseudomonas putida IsoF: The role of quorum sensing
as assessed by proteomics. Sys. App. Microbiol., 28: 87-114.
Augustin M, Ali- Vehmas T, Atroshi F (2004). Assessment of enzymatic
cleaning agents and disinfectants against bacterial biofilms. J.
Pharm. Pharmaceut., 7: 55-64.
Berry D, Xi C, Raskin L (2006). Microbial ecology of drinking water
distribution systems. Cur. Opin. Biotechnol., 17: 297-302.
Bhaskar PV, Bhosle NB (2005). Microbial extracellular polymeric
substances in marine biogeochemical processes. Cur. Sci. 88: 47-53.
Cloete TE (2003). Resistance mechanisms of bacteria to antimicrobial
compounds. Int. Biodeter.Biodegrad., 51: 272-282.
Cloete TE, Jacobs L, Brozel VS (1998). The chemical control of
biofouling in industrial water systems. Biodeter., 9: 23-37.
Czaczyk K, Myszka K (2007). Biosynthesis of extracellular polymeric
substances (EPS) and its role in microbial biofilms formation. Pol. J.
Environ. Stud., 16: 799-806.
de Carvalho CCCR (2007). Biofilms: Recent developments on an old
battle. Patents Biotechnol. 1: 49-57.
Donlan RM (2002). Biofilms: Microbial life on surfaces. Emerg. Infec.
Dis., 8: 1-14.
Flemming HC, Neu TR, Wozniak DJ (2007). EPS matrix: The “House of
Biofilm cells’’. Bacteriol. 189: 7945-7947.
Flemming HC (1998). Relevance of biofilms for the biodeterioration of
surfaces of polymeric materials. Pol. Degrad.,Stab. 59: 309-315.
Frøelund B, Keiding K, Nielsen P (1995). Enzymatic activity in the
activated sludge flocs matrix. App. Microbiol. Biotechnol., 43: 755-
761.
Gaudy AF (1962). Colorimetric determination of protein and
carbohydrate. Indus. Water Waste 7: 17-22.
Gomez – Suarez C, van der Borden PJ, Wingender J, Flemming HC
(2002). Influence of extracellular polymeric susbatnces deposition
and redeposition of Pseudomonas aeruginosa to surface. Microbiol.,
148: 1161-1169.
Johansen C, Falholt P, Gram L (1997). Enzymatic removal and
disinfection of bacterial biofilms. App. Environ. Microbiol., 63: 3724-
3728.
Lapidot A, Romling U, Yaron S (2006). Biofilm formation and the
survival of Salmonella typhimurium on parsley. Int. J. Food Microbiol.,
109: 229-233.
Ledder RG, Timperley AS, Friswell MK, MacFarlane S, McBain AJ
(2008). Coaggregation between and among human intestinal and oral
bacteria. FEMS Microb.Ecol., 66: 630-636.
Lequette Y, Boels G, Clarisse M, Faille C (2010). Using enzymes to
remove biofilms of bacterial isolates samples in the food industry.
Biofoul. 4: 421-431.
Leroy C, Delbarre C, Gillebaert F, Compere C, Combes D (2008). Effect
of commercial enzymes on the adhesion of a marine biofilm forming
bacterium. Biofoul. 24: 11-22.
Liu H, Fang HP (2003). Extraction of extracellular polymeric substances
(EPS) of sludge. Biotechnol. 95: 249-256.
Liu Y, Yang SF, Li Y, Xu H, Qin L, Tay JH (2004). The influence of cell
substratum surface hydrophobicities on microbial attachment.
Biotechnol.110: 251-256.
Loiselle M, Anderson KW (2003). The use of cellulose in inhibiting
biofilms formation from organisms commonly found on medical
implants. Biofoul.19: 77-85.
Melo LF, Bott TR (1997). Biofouling in water systems. Exp. Thermal.
Fluid Sci., 14: 375-381.
O’Toole G, Kaplan HB, Kolter R (2000). Biofilm formation as microbial
development. An. Rev.Microbiol., 54: 49-79.
Orgaz B, Kives J, Pedregosa AM, Monistrol IF, Laborda F, SanJose C
(2006). Bacterial biofilms removal using fungal enzymes. Enz.
Microb. Technol., 40: 51-56.
Parkar SG, Flint SH, Brooks JD (2000). Evalualtion of the effect of
cleaning regimes on biofilms of the thermophillic bacilli on staineless
1524 Afr. J. Microbiol. Res.
steel. App. Microbiol., 96: 110-116.
Pitts B, Hamilton MA, Zelver N, Stewart PS (2003). A microtitter plate
screening method for biofilms disinfection and removal. Microbiol.
Met., 54: 269-276.
Ploux L, Beckendorff S, Nardin M, Neunlist S (2007). Quantitative and
morphological analysis of biofilms formation on self assembled
monolayers. Col. Surf., 57: 174-181.
Prakash B, Veeregowda BM, Krishnappa G (2003). Biofilms: A survival
strategy of bacteria. Cur. Sci., 85: 9-10.
Rochex A, Lebeault JM (2007). Effects of nutrients on biofilms formation
and detachment of Pseudomonas putida strain isolated from a paper
machine. Water Res., 41: 2885-2892.
Simoes M, de Carvalho H, Pereira MO, Viera MJ (2003). Studies on
the behavior of Pseudomonas fluorescens biofilms after Ortho-
phthalaldehyde treatment. Biofoul., 3: 151-157.
Sreenivasan PK, Chorny RC (2005). The effects of disinfectant foam on
microbial biofilms. Biofoul., 21: 141-149.
Stoodley LH, Stoodley P (2002). Developmental regulation of microbial
biofilms. Cur. Op. Biotechnol., 13: 228-233.
Surtherland IW (2001). Biofilm exopolysaccharides: a strong and sticky
framework. Microbiol.147: 3-9.
Ton-That H, Marraffini L, Schneewind O (2004). Protein sorting to the
cell wall envelope of Gram positive bacteria. BBA 1694: 269.
Vickery K, Pajkos A, Cossart Y (2004). Removal of biofilms from
endoscope: Evaluation of detergent efficacy. Inf. Con., 32; 170-176.
Walker SM, Fourgialakis M, Cerezo B, Livens S (2007). Removal of
microbial biofilms from dispense equipment: Effect of enzymatic pre-
digestion and detergent treatment. Inst. Brew., 113: 61-66.
Xavier JB, Picioreanu C, Rani SA, van Loosdrecht MCM, Stewart PS
(2005). Biofilm control strategies based on enzymatic disruption of
the extracellular polymeric substance matrix- a modeling study.
Microbiol., 51: 3817-3832.
Zhang T, Ke SZ, Liu Y, Fang HP (2005). Microbial characteristics of a
methanogenic phenol-degrading sludge. Water Sci. Technol., 52:
73-78.
