Revisiting an agar-based plate method: 
What the static biofilm method can offer 
for biofilm research
Authors: Terhi Oja, Brianna Blomqvist, Kelli Buckingham-
Meyer, Darla Goeres, Pia Vuorela, & Adyary Fallarero
NOTICE: this is the author’s version of a work that was accepted for publication in Journal of 
Microbiological Methods. Changes resulting from the publishing process, such as peer review, 
editing, corrections, structural formatting, and other quality control mechanisms may not be 
reflected in this document. Changes may have been made to this work since it was submitted 
for publication. A definitive version was subsequently published in Journal of Microbiological 
Methods, Volume 107, December 2014, DOI# 10.1016/j.mimet.2014.10.008
Oja, Terhi , Brianna Blomqvist, Kelli Buckingham-Meyer, Darla Goeres, Pia Vuorela, and 
Adyary Fallarero. Revisiting an agar-based plate method: What the static biofilm method 
can offer for biofilm research. Journal of Microbiological Methods. December 2014. Pages 
157-160. https://dx.doi.org/10.1016/j.mimet.2014.10.008
Made available through Montana State University’s ScholarWorks 
scholarworks.montana.edu 
on
th s
me
terial biofilms call for the 
search methods, different to the 
 (Cos et al., 2010). Accordingly, 
ave be
uid she
ctor is
ham-M
and Nelis, 2010).
it is thought to involve several distinctive phases. At the initial phase, 
regularly transferred onto fresh agar medium, while in the static 
biofilm method the biofilm is left to develop on a single filter-covered 
agar plate for the whole incubation period (Fig. 1). The static biofilm 
Consequently, in this contribution we set off to gain a deeper 
 P. aeruginosa ATCC 
ctly according to the 
Meyer et al., 2007). 
ter 1.27 cm, height 0.4 
Revisiting an agar-ba
What the static biofilm
for biofilm 
ce
Bio
euimmersed in liquid, the solid-state growth methods are not as 
common as the well-studied liquid cultures. Among the reported 
cm, BioSurface Technologies Corporation) were placed on an agar 
plate covered with an inoculated filter paper (Whatman Qualitative solid-state methods are the colony biofilm model (Anderl et al., 2000) Grade 2, 70-mm diameter, GE Healthcare). The purpose of the filter by the biofilm matrix. At the final phase, cells return into a planktonic 
stage and detach from the biofilm (Dunne, 2002; Kiedrowski and 
Horswill, 2011; Otto, 2008).
Although bacterial biofilms may also form on surfaces that are not 
Gram-negative Escherichia coli XL-1 Blue and
15442. In all cases, biofilms were grown ex-a
previously described method (Buckingham-
Briefly, sterile borosilicate glass coupons (diameplanktonic bacteria are irreversibly attached to the surface. The 
attachment is followed by aggregation and proliferation of the cells. 
The production and accumulation of extracellular polymeric 
substances then result in the formation of a mature biofilm, where the 
cells are organized into a three-dimensional community surrounded 
under-standing of the development of static method biofilms. For this 
purpose, the kinetics of biofilm development was studied for the 
Gram-positive S. aureus ATCC 25923, a model biofilm-forming 
organism. Furthermore, the results were compared with S. aureus 
ATCC 6538, Staphylococcus epidermidis ATCC 35984, and the and the stat-ic biofilm method (Charaf et al., 
biofilm model, polycarbonate membrane filte
Abbreviations:
 
CFU,
 
ceen studied in detail, and 
on static method plates has not been thoroughly characterized, nor its 
potential as a research choice fully explored.Biofilm formation in liquid cultures has bThe unique properties of bac
development of reliable and specific re
ones optimized for planktonic bacteria
various biofilm culturing methods h
model biofilms grow under various fl
reactors. The choice of the biofilm rea
question under investigation (Buckingen developed, in which 
ar conditions in specific 
 based upon the research 
eyer et al., 2007; Coenye 
method has been shown to be useful in antimicrobial efficacy testing 
against Pseudomonas aeruginosa PAO1, ATCC 15442 and 
Staphylococcus aureus ATCC 6538 biofilms (Buckingham-Meyer et 
al., 2007; Charaf et al., 1999). However, the formation of the biofilms Abstract
The development of biofilms in static plates was m
covered with filter paper, which was inoculated wi
biofilms matrix and biomass were quantified. The 
studies, due to its simplicity and flexibility. 
Terhi Oja & Brianna Blomqvist: Pharmaceutical Scien
University, Turku, Finland
Kelli Buckingham-Meyer & Darla Goeres: Center for 
Bozeman, United States
Pia Vuorela & Adyary Fallarero: Division of Pharmac
University of Helsinki, Finland1999). In the colony 
rs are inoculated and 
olony
 
forming
 
unit;
 
CV,
 
crystal
 
violetitored. Glass coupons were placed on agar 
uspended bacteria. The viable cell density, 
thod is excellent for adhesion and material 
sed plate method: 
 method can offer 
research
s, Department of Biosciences, Abo Akademi 
film Engineering, Montana State University, 
tical Biosciences, CDR, Faculty of Pharmacy, paper is to provide a barrier between the cou-pons and the agar 
surface that allows for diffusion of nutrients while
;
 
TSB,
 
tryptone
 
soy
 
broth;
 
WGA,
 
wheat
 
germ
 
agglutinin.
Fig. 1.Anoverall schemeof themain steps of the static biofilmmethod. The light gray boxes represent TSAor R2A (P. aeruginosa) plates, the gray lines above theplates represent inoculated
filter papers, and the coupons on the filter papers are drawn as empty boxes. The biofilms are drawn in yellow. Only the original 48-h incubation period is shown here.preventing the agar from sticking to the coupon surface. The inoculum
consisted of 1.5 mL of a 1:10 dilution of an 18–24 h old preculture in ei-
ther tryptone soy broth (TSB, Sigma-Aldrich) or in a 100-fold diluted
TSB (only for P. aeruginosa ATCC 15442). After 24 h of incubation in a
humidified incubator at 37 °C, the filters were remoistened with
1.5 mL of 10-fold or 1000-fold (P. aeruginosa ATCC 15442) diluted TSB
(Fig. 1, Buckingham-Meyer et al., 2007).
In this contribution, longer incubation periods for growing the bio-
film were studied (1, 2, 4, 24, 32 or 48 h), and the coupons were ana-
lyzed by either viable cell counts or crystal violet (CV) staining of the
total biomass. In the case of S. aureus ATCC 25923 the couponswere ad-
ditionally analyzed by staining with a wheat germ agglutinin (WGA)
Alexa Fluor 488 conjugate (Molecular Probes, Thermo Fisher Scientific).
At the end of the selected incubation period, each couponwas rinsed by
dipping it in TSB, and subsequently analyzed according to a specific pro-
tocol. Differences in the protocols were only owed to practical reasons,
as the work with three of the strains (S. aureus ATCC 25923, S.
epidermidis ATCC 35984 and E. coli XL-1 Blue) was performed in
Finland while the other strains (S. aureus ATCC 6538 and P. aeruginosa
ATCC 15442) were studied in the United States.
For viable cell counting, the biofilms (except those formedby S. aureus
ATCC 6538 and P. aeruginosaATCC 15442)were removed and dispersed
by a sonication-basedmethod, simpler than the one originally reported
(Buckingham-Meyer et al., 2007; Charaf et al., 1999), which was based
on scraping biofilm off the surface. Coupons were fully immersed in
1 mL of 0.5% (wt./vol.) Tween 20 (Sigma-Aldrich) in TSB, followed by
quick vigorousmixing and 5min of sonication in a water bath sonicator
(Sonorex Digitec, Bandelin) at 25 °C, 35 kHz. In the case of S. aureus
ATCC 6538 and P. aeruginosa ATCC 15442 biofilms, a previously
published protocol was followed (ASTM International, 2013). Briefly:
each coupon was fully immersed in a conical vial containing 10 mL of
buffered water (KH2PO4 0.3 mM, MgCl 6H2O 2.0 mM; pH 7.2). The
vial was vortexed (30 s) and sonicated (30 s, 45 kHz, 25 °C) in 2 cycles
Table 1
The results of the viable counts, CV staining andWGA staining of S. aureusATCC25923 coupons i
inoculumwas increased from1.5 mL to 2.25 mL. TheWGAstainwas quantified atλexcitation = 4
with the standard deviation in parentheses. Samples obtained in each time point were compar
Welch's correction (p b 0.05) (GraphPad Prism program, San Diego, USA).
Time (h) Log (CFU/cm2) CV staining (A
Humidified No humidification Humidified
1 5.55 (±0.24) 5.84 (±0.06) 0.14 (±0.00)
2 5.77 (±0.03) 5.94 (±0.02) 0.14 (±0.01)
4 7.36 (±0.03) 7.38 (±0.22) 0.20 (±0.04)
24 8.23 (±0.62) 7.61 (±0.18) 0.58 (±0.30)
32 8.08 (±0.33) 7.75 (±0.09) 0.85 (±0.09)
48 8.01 (±0.25) 7.49 (±0.30) 1.31 (±0.54)
N.a. = not analyzed.followed by onemore vortexing step for 30 s. In all cases, dilution series
of the resulting suspensions were prepared and plated for the determi-
nation of the colony forming units (CFUs). All plates were incubated at
37 °C for 24 h.
Alternatively, the washed coupons with intact biofilms were ana-
lyzed by staining with CV or the WGA–Alexa fluor conjugate. For all
biofilms except those formed by S. aureus ATCC 6538 and P. aeruginosa
ATCC 15442, the followingmodificationsweremade to a previously de-
scribed CV protocol (Sandberg et al., 2008). The couponswith the intact
biofilm side up were transferred to a 12-well plate, and gently covered
with 70 μL of 2.3% (wt./vol.) CV solution (Sigma-Aldrich) for 5 min.
This volume was enough to cover the surface of the glass coupon
where the biofilm had formed. Washing was done three times with
3 mL of Milli-Q water, and the bound dye was dissolved in 1.5 mL of
96% (vol/vol) ethanol (1 h). In the case of the intact, washed S. aureus
ATCC 6538 and P. aeruginosaATCC 15442 biofilms, the stainingwas per-
formed for 15min in 5mL of 0.5% (wt./vol.) CV (Acros) inMilli-Qwater,
followed by washing three times in 30mL of sterile buffered water. The
dye was dissolved in 5 mL of 95% (vol/vol) ethanol (10 min).
For WGA staining of the S. aureus ATCC 25923 biofilms, a previously
described protocol (Skogman et al., 2012) was followed with minor
modifications: the coupons were washed three times in a 12-well
plate with 3 mL of PBS, and the dye was dissolved in 1 mL of 33% acetic
acid.
The results revealed that for S. aureus ATCC 25923 there were no
significant changes in the viable counts or in the amount of the at-
tached biomass during the first 2 h in a humidified incubator. As
expected, the initial attachment phase was followed by proliferation
of the cells, characterized by a steady increase in the viable counts.
Also, a prominent accumulation phase was observed as a time-shift
in the biomass accumulation: the amount of the biomass attached to
the coupons, measured by CV staining, continued to increase after
24 h, when the number of the CFUs had already reached a maximum
ncubated in a humidified vs. non-humidified incubator. In the latter case, the volume of the
95 nmandλemission = 520 nm. The average value of two biological replicates is presented
ed in humidified vs. non-humidified conditions with an unpaired comparison t-test with
595) WGA staining (fluorescence)
No humidification Humidified No humidification
0.14 (±0.01) 0.23 (±0.05) 0.44 (±0.08)
0.14 (±0.00) 0.46 (±0.25) 0.68 (±0.25)
0.17 (±0.00) 1.89 (±0.25) 2.49 (±0.88)
0.94 (±0.27) 9.68 (±1.31) 9.17 (±2.24)
1.16 (±0.35) N.a. N.a.
1.74 (±0.48) 12.03 (±4.28) 11.83 (±1.77)
59
5
tor.
s an
. Due
ins. Tlevel. The accumulation of the extracellular substances was quantified
by staining with the WGA conjugate that binds to the (poly-)N-
acetylglucosamine residues (PNAG, Sharon, 2007). The PNAG content
0
0,1
0,2
0,3
0,4
0,5
0,6
0,7
0,8
0,9
1
4,5
5
5,5
6
6,5
7
7,5
8
8,5
0 10 20 30 40 50
x
A
59
5
o
lo
g 
(C
FU
/c
m
2 )
time  (h)
S. aureus ATCC 6538
0
0,5
1
1,5
2
2,5
3
4,5
5
5,5
6
6,5
7
7,5
8
8,5
0 10 20 30 40 50
x
A
o
lo
g 
(C
FU
/c
m
2 )
time  (h)
E. coli XL-1 Blue
Fig. 2. The kinetics of biofilm formation for various bacterial species in a humidified incuba
(red circles and red lines). The accumulation of the total biomass on the coupons is shown a
blue lines). The dotted lines were only used to connect the actual experimental data points
and S. aureus 6538 are not directly comparable with the values obtained for the other stracontinued to increase after 24 h, with a similar kinetics to the regis-
tered for the biomass, measured by CV staining. Thus, the CV and
PNAG staining data support that although the cell numbers remained
stable after 24 h, the biomass that the cells were producing continued
to increase. Furthermore, the possibility of using a non-humidified
incubator was also evaluated. The humidification did not cause any
statistically significant change in the kinetics of biofilm development,
suggesting that the higher inoculation volume (2.25 mL) and the
remoistening of the plates likely compensate for the lack of humidifica-
tion (Table 1).
The results indicate that in addition to its use in the analysis of
established biofilms, the static biofilm method could also be useful
in the analysis of the early stages of biofilm formation, e.g. adhesion
studies on various materials. The removal and dispersal of the biofilms
by a simple sonication step enables easy analysis of sample coupons of
varying sizes and shapes. Further flexibility is provided by the choice
of the incubator: established S. aureus biofilms are formed in 48 h
with or without humidification (Table 1).
A major advantage of the static biofilm method is the flexibility,
which includes its usefulness with different bacterial species. A similar
2-h attachment period was observed in all cases, when the kinetics of
biofilm formation were monitored for another S. aureus strain and
three other bacterial species in a humidified incubator. However,
some variation was seen in the length of the subsequent phase: the
highest viable counts were observed at 24 h with E. coli, S. aureus
ATCC 25923 and S. epidermidis, while a 32-h incubation periodwas nec-
essary for the maximal counts of S. aureus ATCC 6538 and P. aeruginosa
ATCC 15442. Interestingly, with E. coli the biomass accumulation
between 24 and 32 h was followed by a significant decline in both the
attached biomass and the viable counts after 32 h (Fig. 2). Thus, it
seems that the length of the incubation period has to be optimized for
each strain.Despite the robustness and the flexibility of the static biofilm meth-
od, there are obvious limitations related to its applicability. Since the
biofilms are formed under no fluid shear, the method can only provide
0
0,1
0,2
0,3
0,4
0,5
0,6
0,7
0,8
0,9
5,5
6
6,5
7
7,5
8
8,5
9
0 10 20 30 40 50
x
A
59
5
o
lo
g 
(C
FU
/c
m
2 )
time  (h)
S. epidermidis ATCC 35984
0
0,1
0,2
0,3
0,4
0,5
0,6
0,7
0,8
0,9
4
4,5
5
5,5
6
6,5
7
7,5
8
0 10 20 30 40 50
x
A
59
5
o
lo
g 
(C
FU
/c
m
2 )
time  (h)
P. aeruginosa ATCC 15442 
The results from the viable counts are presented as average log values of the CFUs per cm2
average absorbance value of the dissolved CV stainmeasured at 595 nm (blue crosses and
to the differences in the staining protocols, the exact values for P. aeruginosa ATCC 15442
he average value and the standard deviation for two biological replicates are presented.useful models of biofilms that are involved in, for instance, the infec-
tions of the ear and the skin. However, there are also numerous benefits
related to the use of themethod. It is simple and economical to use, and
suitable for any laboratory equipped for basicmicrobiologywork,which
makes it a very convenient investigational tool for awider audience. The
method can be directly applied for studies on bacterial adhesion and
biofilm formation on various materials. It is very flexible in terms of
composition, size and shape of the test coupons, and it is currently
used for material testing in our laboratory.
The authors acknowledge the financial support provided by the
Academy of Finland projects WoodyFilm and ArtFilm (decision
numbers 282981 and 272266), as well as by the Drug Discovery and
Chemical Biology network of Biocenter Finland. D. G. was supported
by a Core Fulbright Scholar Grant from the Fulbright Center in Helsinki,
Finland.
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