Received Date : 29-Apr-2016
Accepted Article
Revised Date : 17-Jul-2016
Accepted Date : 05-Aug-2016
Article type
: Original Article
The agar microdilution method – a new method for antimicrobial susceptibility testing
for essential oils and plant extracts
Joanna Golus1, Rafal Sawicki1, Jaroslaw Widelski2, Grazyna Ginalska1
1
2
Department of Biochemistry and Biotechnology, Medical University of Lublin, 1
Chodzki, 20-093 Lublin, Poland
Department of Pharmacognosy with Medicinal Plant Unit, Medical University of Lublin,
1 Chodzki, 20-093 Lublin, Poland
Correspondence
Joanna Golus
Department of Biochemistry and Biotechnology
Medical University of Lublin
1 Chodzki, 20-093 Lublin, Poland.
E-mail address: joanna.golus@umlub.pl
Abbreviated running headline: The agar microdilution method
This article has been accepted for publication and undergone full peer review but has not
been through the copyediting, typesetting, pagination and proofreading process, which may
lead to differences between this version and the Version of Record. Please cite this article as
doi: 10.1111/jam.13253
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ABSTRACT
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Aims
To develop a new agar microdilution technique suitable for the assessment of antimicrobial
activity of natural plant products such as essential oils or plant extracts as well as evaluate the
antimicrobial effect of several essential oils and plant extracts.
Methods and Results
The proposed agar microdilution method was evolved on the basis of the CLSI agar dilution
method, approved for conventional antimicrobials. However, this new method combines
convenience and time/cost effectiveness typical for microtiter methods with the advantages of
the agar dilution of hydrophobic or colouring substances. A different concentrations of the
tested agents were added to eppendorf tubes with molten Mueller-Hinton agar, vortex and
dispensed into the 96-well microplate in a small volume of 100 µl per well which allows for
rapid, easy and economical preparation of samples as well as provides uniform and stable
dispersion without the separation of oil-water phases which occurs in methods with liquid
medium. Next, the agar microdilution plates were inoculated with 4 reference bacterial
strains. The results of our study demonstrated that the minimal inhibitory concentrations
(MICs) were successfully determined using the agar microdilution method even with
hydrophobic essential oils or strongly colouring plant extracts.
Conclusions
The new agar microdilution method avoids the problems associated with testing of water
insoluble, oily or strongly colouring plant natural products. Moreover, it enables reliable,
cheap and easy MIC determination of such agents.
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Significance and Impact of the Study
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In the era of increasing antibiotic resistance high hopes are associated with new drugs of
plant origin. However, the lack of standardized and reliable testing methods for assessing
antibacterial activity of plant natural products causes impediment to research into this area. In
this study we demonstrated that the agar microdilution method can be successfully used for
testing oily and colouring substances.
KEYWORDS
Agar microdilution method
Antibacterial activity
Minimal inhibitory concentration
Essential oil
Plant extract
INTRODUCTION
Increasing drug resistance of microorganisms becomes a serious threat to countering
microbial infections. New, more effective therapies and alternative substances that are
effective against highly resistant strains are still being sought (Alanis 2005; Laxminarayan et
al. 2006; Wise 2011). An alternative to synthetic compounds commonly used in medicine
may be natural substances of plant origin widely distributed in nature (Cowan 1999; Carson
et al. 2006; Bakkali et al. 2008; Aleksic and Knezevic 2014). However, there is no standard
for assessing antibacterial activity of plant natural products such as essential oils (EOs) or
plant extracts. The lack of standardized and reliable testing methods causes impediment to
research into the antimicrobial activity of these products. The generally applicable Clinical
and Laboratory Standards Institute methods (CLSI 2015) are standardized only for
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conventional antimicrobial agents such as antibiotics. Modifications of these methods are
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implemented in the examination of other molecules, such as EOs. Nevertheless, the obtained
results vary between publications due to differences in research methodologies so comparing
the data from different studies is problematic. The necessity of developing a standard and
reproducible technique for assessing plant oils and extracts has been highlighted by numerous
authors (Mann and Markham 1998; Hammer et al. 1999; Cavanagh and Wilkinson 2002;
Lahlou 2004; Ncube et al. 2008; Das et al. 2010; Horváth and Ács 2015; Tan and Lim 2015).
The greatest difficulties connected with testing EOs result from their volatility, water
insolubility and complexity. The hydrophobic nature of EOs and their high viscosity may
reduce the dilution capability and cause unequal distribution of the oil through the medium.
On the other hand, plant extracts are often strongly colouring and also poorly soluble in the
medium (Mann and Markham 1998; Kalemba and Kunicka 2003; Tan and Lim 2015).
The most popular techniques used for the assessment of antimicrobial activities of
plant oils and extracts are the agar diffusion method (disc diffusion or well diffusion) and the
dilution method (agar dilution and broth macro/microdilution) (Kalemba and Kunicka 2003;
Tan and Lim 2015). However, the usefulness of the agar diffusion techniques is limited to the
generation of only preliminary, qualitative data since the poorly soluble components of
essential oils and plant extracts do not diffuse well in the agar medium. Moreover, these
methods are considered to be inappropriate for EOs as their volatile components are likely to
evaporate during the incubation time (Mann and Markham 1998; Hammer et al. 1999;
Kalemba and Kunicka 2003; Ncube et al. 2008; Tan and Lim 2015).
In contrast to diffusion methods, the dilution methods allow quantitative assessment
of the antimicrobial susceptibility by determining the lowest concentration of the agent
capable of inhibiting the growth of the tested organism, which is described as minimum
inhibitory concentration (MIC). In the case of the broth dilutions, both macro- and
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microdilution methods are similar and well-established. Nonetheless, due to the use of
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smaller amounts of medium, reagents, and tested agents the microdiluton method is more
economical and less laborious than the macrodilution method. Thus, the miniaturization made
the microdilution method more practical and popular in testing conventional antimicrobial
agents (Wiegand et al. 2008; Jorgensen and Ferraro 2009).
However, the use of this method for assessing the antimicrobial activity of plant oils
and extracts entails many difficulties and is often associated with misinterpretation of results.
As a lot of authors noted, the main reason for the difficulties of using the broth microdilution
is the problem with dispersion of water insoluble compounds in the liquid growth medium
(Mann and Markham 1998; Lahlou 2004; Tan and Lim 2015). Despite the use of dispersing
and emulsifying agents the hydrophobic oily substances are often poorly soluble in the liquid
medium and the separation of oil-water phases occurs. In such circumstances, even contact
between the test organism and the agent is not ensured. Furthermore, the determination of
MIC value becomes problematic when the opacity of oil-water emulsions interferes with the
turbidity of bacterial growth (Mann and Markham 1998; Carson et al. 2006; Chorianopoulos
et al. 2006; Tan and Lim 2015). Similarly, strongly colouring compounds or extracts, also
make the MIC value impossible to be determined by broth microdilution because they
preclude distinction between bacterial growth and the medium (Tan and Lim 2015). In such
conditions, even applying growth indicators, as some authors suggest (Mann and Markham
1998; Rahman et al. 2003; Ncube et al. 2008), could be useless because of the difficulty in
determining an indicator colour change in strongly coloured or high opaque medium.
Due to these difficulties, a technique that is more optimal for assessing antibacterial
activity of plant extracts and oils is the agar dilution method (Silva et al. 2005; Tan and Lim
2015). The main advantage of this method is the provision of uniform and stable dispersion
of the oils and extracts when they are incorporated into the agar medium (Santos et al. 1997;
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Silva et al. 2005). It has been demonstrated (Remmal et al. 1993; Mann and Markham 1998;
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Lahlou 2004) that the addition of merely 0.2% agar as a stabilizer overcomes the problem of
emulsion stability for essential oils in liquid medium. Another advantage of the agar dilution
method results from the bacteria’s ability to form visible growth on the solid agar medium
which could be easily detected by the unaided eye. Thereby, all the changes in opacity or
colour of the solid medium under the influence of tested agents become irrelevant to detect
bacterial growth. So, in the case of strongly colouring or opaque medium, the determination
of bacterial growth on the surface of agar is simpler and clearer than assessment of turbidity
change by broth dilution technique. Nevertheless, the agar dilution method is not popular in
the context of essential oils and plant extracts because it requires large amounts of tested
agents in preparation of agar plates, each containing a different concentration of the agent
(Tan and Lim 2015). It is a limitation especially when the examined antimicrobial agents are
expensive or obtained in a microgramme scale. Moreover, this technique is both tedious and
labour-intensive as well as it requires the relatively large amount of materials and space for
each test (Wiegand et al. 2008; Tan and Lim 2015). To overcome all of the above mentioned
problems, we propose the miniaturization of the standardized agar dilution method to adapt
them for convenient testing of hydrophobic, oil-based or strongly colouring molecules.
The aim of this study was to develop an agar microdilution technique suitable for the
assessment of antimicrobial activity of natural plant products such as essential oils or plant
extracts as well as evaluate the antimicrobial effect of 17 commercial essential oils and 4 new
plant extracts.
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MATERIALS AND METHODS
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Tested organisms and growth conditions
The reference bacterial strains used in this study were Staphylococcus aureus ATCC
25923, Staph. epidermidis ATCC 12228, Escherichia coli ATCC 25922 and Pseudomonas
aeruginosa ATCC 27853. All strains were obtained from the American Type Culture
Collection. Stock cultures were stored at -70°C in Viabank™ vials (Medical Wire &
Equipment, England). Intermediate cultures were prepared from the stock cultures and used
to create fresh 24-hour cultures before each experiment. All cultures were grown at 35°C on
the Mueller-Hinton agar (Oxoid, United Kingdom).
Reference antimicrobials
Gentamicin, tetracycline, ciprofloxacin and ceftriaxone (Sigma-Aldrich, USA) were
used as reference standard. Stock solutions were prepared according to the manufacturer’s
instruction. Intermediate (10× concentrated) solutions were prepared by making serial
twofold dilutions in Mueller-Hinton broth (Oxoid, United Kingdom) in the range from 64 µg
ml-1 to 0.008 µg ml-1.
Essential oils
The essential oils used in this study were obtained from commercial suppliers. They
were as follows cinnamon bark oil (Unimark Remedies Ltd., India), bergamot oil, clove oil,
coriander oil, juniper oil, lavender oil, pine oil (Pollena – Aroma, Poland), basil oil,
cedarwood oil, lemon balm oil, marjoram oil, rosemary oil (Tag-Pol J.V./Sabana Oil,
Poland), eucalyptus oil, fennel oil, mint oil (Kej, Poland), hyssop oil (Vera, Poland) and tea
tree oil (Melaleuca Poland, Poland). All the EOs were 100% pure and tested in the
concentration range from 8 to 0.004% (v/v). Serial twofold dilutions of the EOs tested in the
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range from 1 to 0.004% (v/v) were made in dimethyl sulfoxide (DMSO, Sigma-Aldrich,
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USA). Next, the appropriate amount of molten Mueller-Hinton agar (Oxoid, United
Kingdom) was added. The final concentration of DMSO in the agar per well did not exceed
1% (v/v). In the case of higher concentrations of the EOs (8 – 2% v/v) it was not possible to
make serial twofold dilutions in DMSO and not exceed 2% of DMSO in the final
concentration in agar. So the required volumes of EOs were directly dispensed to eppendorf
tubes and DMSO were added in an amount equal to 2% final concentration per well. The
tubes were vortexed and then the appropriate amount of molten Mueller-Hinton agar was
added. All the tubes were vortexed and kept at 50°C in ThermoMixer as described in the agar
microdilution procedure.
Plant extracts
Seseli devenyense Simonkai, Portenschlagiella ramosissima Tutin and Peucedanum
luxurians Tamam belonging to the Apiaceae family were used in this study. The mature fruits
of S. devenyense and P. ramosissima were collected from the Pharmacognostic Garden of the
Department of Pharmacognosy with Medicinal Plant Unit (Medical University of Lublin,
Poland). The mature herbs and fruit of P. luxurians were collected from the Botanical Garden
of the Adam Mickiewicz University in Poznań (Poland). The collected plant materials were
air dried, pulverized using a mill and immediately subjected to extraction. Samples (50 g)
were extracted with hot, pure methanol (3 x 500 ml each time for 30 minutes) on a water bath
at the temperature of 70ºC. Extracts from the same sample were combined and evaporated to
the dryness under reduced pressure at 50ºC. The extracts were tested in the concentration
range from 40 to 0.06 mg ml-1. Intermediate (10× concentrated) solutions were prepared by
making serial twofold dilutions in Mueller-Hinton broth.
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Inoculum preparation
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Inoculum was prepared according to CLSI direct colony suspension method. Briefly,
fresh (18- to 24-hour) bacterial colonies were suspended in Mueller-Hinton broth (Oxoid,
United Kingdom) to achieve a turbidity of 0.5 McFarland standard corresponding to
approximately 1 to 2 × 108 colony-forming units (CFU) ml-1. Turbidity of bacterial
suspensions was measured using a nephelometer (PhoenixSpec Nephelometer, Becton
Dickinson, USA). Next, 0.5 McFarland suspensions were diluted 1:10 in fresh MuellerHinton broth to obtain a concentration of 107 CFU ml-1 and than 2 µl were applied to the agar
to each well of microplate, to give a final inoculum density of approximately 104 CFU per
spot. Bacterial suspensions were used within 15 minutes after preparation.
Agar microdilution procedure
The agar dilution method approved by the CLSI (2015) has been modified to adapt it
for testing very small amounts of essential oils and plant extracts. Assays were performed in
reduced volumes of 100 µl on sterile 96-well microplates with round bottom or optionally at
200 µl on microplates with flat bottom (Anicrin, Italy).
Previously prepared intermediate (10× concentrated) solutions of the tested molecules
(10 µl per each replicate) were added to eppendorf tubes with molten Mueller-Hinton agar
(90 µl per each replicate). The tubes were vortexed and kept at 50°C in ThermoMixer (HLC –
M1R 23, DITABIS, Germany) until they had been dispensed into the 96-well microplate (100
µl per well). Each twofold serial dilution was tested in triplicate. The final concentration of
DMSO (1 or 2%) used in this study had no effect on the growth of tested bacteria. At the
same microplate the sterility control, the growth control and the control for DMSO
(respectively 1 or 2%) were carried out for each tested strain. The plate was kept at room
temperature until the agar had solidified. When the agar surface was dry the microplate was
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inoculated with 2 µl of freshly prepared inoculum using a multichannel micropipette or the
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multiple dispensing mode of the electronic pipette. The inoculum was prepared as described
above so that the final concentration was 104 CFU per spot. The inoculated plates were
incubated at room temperature until the inoculum had been absorbed unto the agar. Next, the
plates were sealed in a plastic bag or with a sealing film (Parafilm®, Sigma-Aldrich, USA) to
prevent drying. Additionally, the corner wells were omitted and filled with sterile water. This
wells evaporate the fastest which could influence the bacterial growth. The plates were
incubated at 35°C for 16 to 20 hours.
The results were determined on a dark and nonreflecting surface. The MIC was
recorded as the lowest concentration of the tested agent that completely inhibits bacterial
growth. According to the CLSI recommendations the presence of a single colony or a faint
haze caused by the inoculum was disregarded. Each experiment was repeated three times.
RESULTS
On the basis of MIC values obtained for reference antimicrobials (Table 1) the
susceptibility and resistance interpretations were designated. E. coli ATCC 25922 and S.
aureus ATCC 25923 were susceptible to all the tested reference antimicrobials (ceftriaxone,
ciprofloxacin, gentamicin and tetracycline) while the tetracycline MIC determined for S.
epidermidis ATCC 12228 was in the resistance range. In the case of P. aeruginosa ATCC
27853 it was interpreted as susceptible to ciprofloxacin and gentamicin and resistant to
ceftriaxone and tetracycline. The results confirmed that the MIC values estimated by the new
agar microdilution method are in accordance with the CLSI Quality Control Ranges for all
the tested reference antimicrobials (CLSI 2016).
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The results of our study demonstrated that even the strong colour and turbidity of
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medium obtained after preparing the desired concentrations of plant extracts and EOs did not
prevent the proper determination of MIC. Because the assays were performed in small
volumes, agar with tested molecules dispensed into the microplate wells solidified quickly,
which did not lead to the separation of oil-water phases. The obtained dispersions were
homogeneous and stable, but they were considerably different in transparency and colour.
The higher the concentration of oil or extract was tested, the more opaque and coloured the
medium was. In order to illustrate that differences we have prepared the example of the agar
microplate which is shown in Fig. 1. We have presented each twofold dilution of a particular
agent in two replications to allow the comparison of the inoculated wells (columns 2, 4, 6, 8,
10, 12) with the uninoculated control wells (columns 1, 3, 5, 7, 9, 11). The use of gentamicin
(Fig. 1b, columns 1, 2) displays how the plant origin samples could be different from
standard antimicrobial agents. The MIC of each tested agent can be seen as the first well of
each inoculated column showing no bacterial growth. The examples of strongly colouring
and high opaque samples are presented in the Fig.1 P. ramosissima extract (Fig. 1b, columns
3 and 4) and P. luxurians extract (Fig. 1b, columns 5 and 6). In spite of their specificity, the
bacterial growth on the agar surface is clearly visible (Fig. 1b, columns 4 and 6) and easily
distinguishable from uninoculated wells (Fig. 1b, columns 3 and 5). In the case of essential
oils samples presented in Fig. 1 two of them are extremely opaque, especially in high
concentrations. Despite this, the MIC values are easy to be determined as follow: fennel oil
MIC > 8% (Fig. 1b, column 8), cedarwood oil MIC 2% (Fig. 1b, column 10) and cinnamon
oil MIC 0.015% (Fig. 1, column 12) against reference strain of Staph. aureus ATCC 25923.
The antibacterial activity of the 17 commercial essential oils against 4 reference
bacterial strains estimated by the agar microdilution method are listed in Table 2. The
obtained MICs were within a broad range from 0.015% to 8% depending on the tested oil and
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strain. A total of 11 of the studied EOs inhibited growth of all the tested strains. Both Staph.
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aureus and Staph. epidermidis were inhibited by all the tested oils apart from fennel oil,
while E. coli was inhibited by all the oils except cedarwood one. Ps. aeruginosa was
inhibited by 11 out of 17 tested oils and in comparison with other strains, higher
concentrations of oils were needed to inhibit its growth. The greatest antimicrobial activity
against all the tested strains was detected to cinnamon oil with MIC of 0.015% against Staph.
aureus and Staph. epidermidis, 0.03% against E. coli and 0.06% against Ps. aeruginosa. Also
high antibacterial activity was found for pine oil, clove oil and tea tree oil with MIC of
0.03%, 0.12%, 0.06% respectively against Staph. aureus; 0.03%, 0.06%, 0.06% against
Staph. epidermidis; 0.06%, 0.06%, 0.12% against E. coli and 0.25%, 1%, 0.25% against Ps.
aeruginosa.
Regarding the examined plant extracts, they did not demonstrate a high antibacterial
activity (Table 3), though all the four extracts were active against Staph. aureus and Staph.
epidermidis and two of them against Ps. aeruginosa. No inhibition of growth was observed in
the case of E. coli even at the highest concentration of 40 mg ml-1.
DISCUSSION
Natural substances of plant origin may successfully constitute an alternative to
synthetic compounds commonly used in medicine as well as supplement conventional
therapies. They are widely prevalent in nature and many of them have been used for centuries
in traditional medicine (Carson et al. 2006; Fadli et al. 2012; Aleksic et al. 2014; Horváth
and Ács 2015). However, the research on the antimicrobial activity of natural plant products
has been hindered by the deficiency of standardized and reliable testing methods. Various
publications have presented the antimicrobial activity of essential oils and plant extracts;
nonetheless, large differences in MIC values are obtained by different authors and the results
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are not directly comparable due to methodological dissimilarities (Mann and Markham 1998;
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Hammer et al. 1999; Cavanagh and Wilkinson 2002; Lahlou 2004; Tan and Lim 2015).
To solve the problem of the selection of an appropriate method for testing oil-based or
strongly colouring plant oils and extracts, we have proposed the miniaturization of the
standard agar dilution method. The suggested agar microdilution method is based on the
CLSI recommended standards (CLSI 2015) but it enables to test very small amounts of
potential antimicrobials compared with standard agar dilution. The amount of 20 ml of the
tested agent solution is necessary to evaluate merely a single dilution in the standard agar
dilution procedure, whereas only 0.1 ml is required for a single dilution of the tested agent in
our agar microdilution procedure. As a general principle, each antimicrobial agent is
examined in serial twofold dilution and in repetitions, therefore very large amounts of tested
substances are required in the standard method. The method we propose allows to reduce the
quantity of studied substances 200 times. Also, great savings of disposable laboratory
materials, reagents and space are worth mentioning. The 96-well microplate used in the agar
microdilution enables to test 12 substances in a range of 8 two-fold dilutions which would
require the preparation of a stack of 96 Petri dishes in the case of standard agar dilution.
When expensive substances are used, the possibility of using a small quantity is of great
significance. Moreover, the method we propose is considerably simpler and substantially less
cumbersome than the agar dilution method. The small volume allows for rapid and easy
preparation of homogeneous solutions of tested agents. Thereby, simultaneous work with a
great number of molecules at different concentrations could be easily manageable.
Because the agar microdilution assays were performed in small volumes, agar with
tested molecules dispensed into the microplate wells solidified quickly which did not lead to
the separation of oil-water phases as it happens in the liquid growth media. Our unpublished
data revealed that the MIC values obtained for the same EOs by the broth microdilution
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method were higher than these obtained by the agar microdilution. Thus, the separation of
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oil-water phases possibly gives bacteria a greater chance of survival and growth, thereby
overstating the obtained MIC values. It could explain why some authors receive a
fundamental difference for some oils when comparing the MICs obtained by agar and broth
methods. Hammer et al. (1999) compared the MICs of 20 EOs and stated the substantial
differences in MICs, respectively, 0.06% (agar) and > 8% (broth) for sandalwood oil, 0.12%
and > 4% for vetiver oil, 0.5% and > 2% for patchouli oil against Candida albicans. Such
differences are much greater than the acceptable ones in terms of one twofold dilution and
undoubtedly they are caused by the specificity of the EOs.
The advantage of the standard agar dilution method is the possibility of testing a large
number of bacterial strains simultaneously onto one agar plate. However, this method is not
the method of choice if susceptibility to a broad range of different substances is to be tested
on a smaller number of bacterial strains (Wiegand et al. 2008). The agar dilution and broth
macrodilution methods which require large volumes of tested substances are not
recommended for most routine procedures (Wiegand et al. 2008; Jorgensen and Ferraro
2009; Tan and Lim 2015). Therefore, the agar microdilution method offers a useful
alternative to current standard methods.
Visual assessment of bacterial growth inhibition on the agar surface is definitely easy
to be carried out and consistent with the CLSI standards (CLSI 2015). No additional steps are
taken as it occurs in the case of broth dilution methods which employ growth indicators
(Mann and Markham 1998; Rahman et al. 2003; Ncube et al. 2008). Moreover, in methods
which need the use of a growth indicator, it could be necessary to assess reduction of
densities for each tested organism (Mann and Markham 1998), which additionally
complicates the procedure. The method we suggest requires neither additional steps after
incubation nor special adaptation of a test for the particular organism.
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The essential oils and plant extracts used in this study have been chosen to create a
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diverse panel of test substances which are known as very troublesome for antibacterial
susceptibility testing. Our results confirm that even strongly colouring extracts as well as
poorly soluble oils incorporated into the agar medium gave the uniform distribution of the
tested molecules and then it was proved that MIC values had been reliably predicted using
the agar microdilution. The EOs tested at various concentrations were significantly different
in transparency. In spite of the fact that the oils studied at high concentrations were the most
opaque, the determination of MIC values did not pose a problem. In the case of the examined
plant extracts we demonstrated that even the strong colour of medium obtained after
preparing the desired concentrations of extracts did not prevent the proper determination of
MIC.
To summarize the MIC values obtained for the EOs, the highest activity was detected
to cinnamon bark oil, with the lowest MIC of 0,015% against Staph. aureus and Staph.
epidermidis. Other oils with high antibacterial activity against all the tested strains include
pine, tea tree, clove and rosemary ones. Regarding the examined plant extracts, they
exhibited moderate antibacterial activity against three of the tested strains and no activity
against E. coli, but we demonstrated that even such a difficult material could be successfully
analysed by means of our method.
The research showed that some of the tested essential oils have a great potential to
control bacterial infections. Conducting a subsequent study with selected oils against clinical
bacterial strains characterized by a high degree of antibiotic resistance can provide a variety
of clinically significant information. It also seems important to examine the exact chemical
composition of the highest activity essential oils and to test the antimicrobial activity of these
components. Such knowledge could allow for effective industrial application of plant origin
antimicrobials.
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In conclusion, the agar microdilution method successfully avoids the hitherto
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encountered problems of testing oily or strongly colouring molecules and allows reliable
determination of MICs for essential oils and plant extracts irrespective of their colour or
opacity. This new method combines convenience and time/cost effectiveness typical for
microtiter methods with the advantages of the agar dilution of hydrophobic or colouring
substances. Due to the fact that the method we propose is performed in accordance with the
CLSI standards in terms of inoculum preparation, serial twofold dilutions of tested agents and
the way of determination of the results, the agar microdilution enables direct comparisons of
the results and global application of the miniaturization method. Therefore, this cheap and
efficient method constitutes a helpful and reliable tool for working with oily and colouring
agents and can be easily performed in any microbiological laboratory.
ACKNOWLEDGMENTS
Financial assistance was provided by the Ministry of Science and Higher Education in
Poland within the DS2/15 project of the Medical University of Lublin. This work was
developed using the equipment purchased within the agreement No. POPW.01.03.00-06010/09-00 Operational Program Development of Eastern Poland 2007-2013, Priority Axis I,
Modern Economy, Operations 1.3. Innovations Promotion.
CONFLICT OF INTEREST STATEMENT
The authors declare no conflict of interest.
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TABLE
Table 1 MICs of reference antimicrobials (µg ml-1) determined by the agar microdilution
method
Antimicrobials
Staphylococcus
aureus
ATCC 25923
Staphylococcus
epidermidis
ATCC 12228
Escherichia
coli
ATCC 25922
Pseudomonas
aeruginosa
ATCC 27853
Ceftriaxone
Ciprofloxacin
Gentamicin
Tetracycline
2
0.5
0.25
0.25
1
0.12
0.12
32
0.06
0.015
0.5
1
32
0.5
2
32
Table 2 MICs of 17 essential oils (% v/v) determined by the agar microdilution method
Essential oil
Basil oil
Bergamot oil
Cedarwood oil
Cinnamon bark oil
Clove oil
Coriander oil
Eucalyptus oil
Fennel oil
Hyssop oil
Juniper oil
Lavender oil
Test organism
Staphylococcus Staphylococcus
aureus
epidermidis
ATCC 25923
ATCC 12228
1
4
2
0.015
0.12
0.5
1
>8
2
0.06
2
0,5
4
4
0.015
0.06
0.5
0.5
>8
2
0.12
4
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Escherichia
coli
ATCC 25922
Pseudomonas
aeruginosa
ATCC 27853
2
4
>8
0.03
0.06
0.5
2
8
1
0.25
1
4
>8
>8
0.06
1
4
2
>8
4
4
>8
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Lemon balm oil
Marjoram oil
Mint oil
Pine oil
Rosemary oil
Tea tree oil
0.25
1
0.5
0.03
0.06
0.06
0.25
2
0.5
0.03
0.12
0.06
0.5
0.5
0.5
0.06
0.25
0.12
>8
8
>8
0.25
1
0.25
The highest tested oil concentration was 8% (v/v).
MICs of oils that did not show antimicrobial activity in the range tested are listed as “> 8”.
Table 3 MICs of plant extracts (mg ml-1) determined by the agar microdilution method
Plant Extract
P. luxurians - herbs
P. luxurians - fruit
P. ramosissima - fruit
S. devenyense - fruit
Staphylococcus Staphylococcus
aureus
epidermidis
ATCC 25923
ATCC 12228
10
20
40
20
5
20
20
20
Escherichia
coli
ATCC 25922
Pseudomonas
aeruginosa
ATCC 27853
> 40
> 40
> 40
> 40
> 40
40
> 40
40
The highest tested extract concentration was 40 mg/ml.
MICs of extracts that did not show antimicrobial activity in the range tested are listed as “> 40”.
FIGURE LEGEND
Figure 1 The agar microdilution test
(a) Outline of the setup of the agar microdilution plate; + inoculum – the inoculated
columns; SC – sterility control; GC – growth control;
(b) Dark background photography of the 96-well microplate with serial twofold dilutions of
the tested molecules in agar, after 20 h of incubation at 35°C with Staph. aureus ATCC
25923 inoculum.
(c) The magnifications of selected wells (white frames) – the inoculated wells compared with
the uninoculated control wells on the example of: (I) the sterility and growth controls,
(II) P. ramosissima fruit extract, (III) fennel oil.
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