Food Control 22 (2011) 360e368
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Food Control
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Control of food spoilage fungi by ethanol
Thien Dao, Philippe Dantigny*
Laboratoire de Génie des Procédés Microbiologiques et Alimentaires, Université de Bourgogne, Agro-Sup Dijon, France
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 11 March 2010
Received in revised form
2 August 2010
Accepted 10 September 2010
This review discusses the effects of ethanol on the inhibition of growth and germination of fungi and on
the inactivation of fungal spores. After a brief survey on the impact of spoilage fungi on the economy and
food quality, the major applications of ethanol in controlling fruit decay and extending the shelf-life of
food products are reviewed. Many parameters including minimum inhibitory concentration (MIC) and
D-values for various moulds are included. The thermodynamic relationship between the liquid phase and
the headspace and the mode of action of ethanol on fungi are explained. Due to their promising use as
a fumigant, special attention is paid to ethanol vapours.
Ó 2010 Elsevier Ltd. All rights reserved.
Keywords:
Ethanol
Conidia
Fungi
Mould
Inhibition
Inactivation
Predictive mycology
1. Introduction
Moulds can grow on a great variety of substrates and a wide
range of pH, water activity (aw) and temperature. Fruits juices are
usually quite acid, in the range pH 1.8e2.2 (lemons) to 4.5e5.0
(tomatoes), but the rind tissue that fungal postharvest pathogens
colonize is about pH 5 to 5.5 (lemons). Tissue pH alone does not
account for the dominance of fungi as plant pathogens. Fungi are
capable of rapid invasive growth into tissue, they have developed
environmental sensing mechanisms, enabling them to tailor in
ambient conditions, by acidification and alkalinisation, to best fit
their offensive arsenal (Prusky & Yakoby, 2003). Fungi are capable
of rapid invasive growth into tissue while bacteria are not. Therefore microbial spoilage of fruit and fruit products is always caused
by fungi (Pitt & Hocking, 1999). Most Penicillia can develop as low
as pH 2 (Panasenko, 1967) and many are postharvest pathogens.
Almost all xerophilic fungi (i.e., capable of growth at 0.75 aw) are
ascomycetes that produce ascospores in asci. Amongst the ascomycetes, Eurotium species are the most common causes of spoilage
of dried cereals. Other ascomycetes (e.g., Byssochlamys species,
Eupenicillium lapidosum, Neosartorya fischeri, Talaromyces species)
are producing ascospores of very high heat resistance which can
* Corresponding author. Laboratoire de Génie des Procédés Microbiologiques et
Alimentaires, AgroSup Dijon, 1 Esplanade Erasme, 21000 Dijon, France. Tel.: þ33 (0)
3 80 70 44 71.
E-mail address: [email protected] (P. Dantigny).
0956-7135/$ e see front matter Ó 2010 Elsevier Ltd. All rights reserved.
doi:10.1016/j.foodcont.2010.09.019
survive heat processing and are responsible for spoilage of
pasteurized foods. In addition, most of the moulds with the notable
exception of the Aspergilli exhibited a minimum temperature for
growth close to the freezing point.
Moulds can develop at the field level. It has been reported that
25% of agriculture products are contaminated with mycotoxins
(Mannon & Jonhson, 1985). But moulds can also develop during
storage of raw products, and subsequent transport and sale causing
considerable economic losses annually for food manufacturers and
consumers alike. It is very difficult to assess losses attributable to
moulds. In the baking industry, these losses varied between 1% and
3% of products depending on season, type of product and method of
processing (Malkki & Rauha, 1978). Another estimate from one
bakery in the US was 5% losses (Killian & Krueger, 1983). Even
assuming only 1% losses, moulds could be spoiling over 23,000 tons
of bread worth nearly £20 million in the UK every year. Throughout
Western Europe the annual losses could be around 225,000
tons of bread worth £242 million (Legan, 1993). More generally
losses of food to fungal spoilage in Australia must be in excess of
$10,000,000 per annum: losses in damp tropical climate and
countries with less developed technology remain staggering (Pitt &
Hocking, 1999). In the fruit industry, postharvest losses are 5e10%
when postharvest fungicides are used (Cappellini & Ceponis, 1984).
Without fungicides, losses of 50% or higher have occurred in some
years. For example, in a 1993 test to assess the decay potential of
stone fruit, an average of 52.8% (range 15e100%) of the fruit
decayed during the ripening of eight collections that had not been
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
treated with postharvest fungicides (Margosan, Smilanick,
Simmons, & Henson, 1997).
Moulds are disseminated through spores that are produced in
the environment. Air can be a vector in the distribution of almost all
fungi that are relevant to food spoilage but packaging materials
may also be source of fungal contamination (Scholte, 1995). It is
troublesome that after germination of the spores, moulds spread
rapidly by aerial mycelia along the fruits, cereals, and food. In order
to delay mould growth and subsequent production of metabolites,
amongst which the most important for food quality and safety are
mycotoxins (D’Mello & MacDonald, 1997) it would be highly
desirable to control food spoilage fungi.
Among the physicochemical hurdles cited in the literature,
ethanol is recognized as a mould inhibitor (Legan, 1993). For a long
time, ethanol has been used as a fungicide treatment. Ethanol is used
commercially in a lot of products, such as perfumes, many food
products paints, alcoholic beverages and additives. Ethanol is a small
molecule produced either by chemical synthesis or by microbial
fermentation. Recently, there has been substantial interest in nonbiological control agents as well as biological control agents to
replace the existing chemical applications. Non-biological control
involves chemicals that are Generally Regarded as Safe “GRAS”
product such as ethanol (Karabulut, Mlikota Gabler, Mansour, &
Smilanick, 2004; Romanazzi, Karabulut, & Smilanick, 2007) as
alternative treatments. As such it can be used in the food industry.
Ethanol is also easily miscible with water in all proportions. It is
a good solvent, as starting compound for the manufacture of dyes,
cosmetic and explosives. Ethanol is a familiar constituent of many
beverages and is considered to be the least toxic of the straight-chain
alcohols (methanol, ethanol, propanol, butanol, etc) evaporates
quickly at room temperature, not leaving any residues (Nittérus,
2000). For a long time, it is also as a disinfectant due to its antibacterial properties. Ethanol is a common food component with potent
antimicrobial activity (Feliciano, Feliciano, Vendruscuolo, Adaskaveg,
& Ogawa, 1992; Karabulut, Smilanick, Mlikota Gabler, Mansour, &
Droby, 2003; Larson & Morton, 1991). Smith (1947) concluded that
95% ethanol was best for wet surfaces, 50% for dry surfaces, and 70%
for either wet or dry surfaces. This percentage has been shown to be
a superior surface disinfectant properties to control bacteria in hand
washing, skin de-germing, or instrument sanitation tests (Ali, Dolan,
Fendler, & Larson, 2001). But there is no evidence that this ethanol
concentration is also optimum to kill fungi.
The inhibitory effect can be obtained by adding ethanol directly
to the product or by using ethanol vapours. Many studies were
concerned with the use of ethanol for extending the shelf-life of
food products and for reducing fruit decay. In a first section, these
studies will be reviewed. Although this literature did provide useful
guidelines for preserving food and agriculture products most of the
previous studies were concerned with natural microflora, including
bacteria, yeasts and moulds. Depending on its concentration, on the
type of product and organisms, ethanol may have different effects
on mould development. In a second section, the inhibitory effects of
ethanol on growth and germination and on the inactivation of
spores will be reviewed. In contrast to the previous section, these
studies were concerned with some moulds isolated from spoiled
products and cultivated in pure cultures. The effects of ethanol
(either through liquid application or vapours in the headspace) will
be assessed under controlled conditions in terms of ethanol
concentration, temperature, pH and water activity. In fact, it was
pointed out that when ethanol vapour generators were used for
preserving foods, the concentration of ethanol in the headspace
was not constant (Daifas, Smith, Tarte, Blanchfield, & Austin, 2000;
Daifas et al., 2003).
In the third section the application of liquid ethanol will be
compared to the use of ethanol vapours in terms of origin of the
361
contaminations, regulations and efficacy of the treatments. There is
much controversy about the mode of action of ethanol. Seiler
(1989) found no difference in the inhibitory effect of ethanol
applied as liquid or as vapour. In contrast, Smith et al. (1987)
reported that lower levels of ethanol were required in the vapour
phase compared to direct addition to a medium or food for
complete microbial inhibition. According to Lerici and Manzocco
(2000), the toxicity of ethanol can be described by its ethanol
vapour pressure. This assumption is in accordance with the
increase of ethanol toxicity with temperature. However, recent
studies (Dantigny, Dao, Dejardin, & Bensoussan, 2007) have shown
that the toxicity of ethanol cannot be explained by its vapour
pressure only. The mode of action of ethanol should also be evaluated at the cell level. The fourth section aimed at reviewing the
biological principles for the effects of ethanol on fungi.
2. Industrial use of ethanol
2.1. Extension of mould free shelf-life
Addition of ethanol increases the mould free shelf-life of bread
when added after baking and cooling at concentrations from 0.5 to
3.5% (wt/wt) of loaf weight (Geiges & Kuchen, 1981; Plemons, Staff
& Cameron, 1976; Seiler, 1984; Seiler & Russell, 1991; Vora & Sidhu,
1987). Differences in methods between workers make it difficult to
compare results. The data of greatest practical use are those of
Seiler (1984), which show that the shelf-life increases with ethanol
concentration, a 50% extension in life being obtained with the
addition of 0.5% ethanol based on loaf weight.
Ethanol is only permitted at levels up to 2% by product weight in
pizza whether the alcohol is added directly to the food or not. In
Italy, sandwich loaves named “pancarrè per tramezzini” contain up
to 2% ethanol wt/dry wt and do not need any addition of sorbic or
propionic acid (Bonetto & Bortoli, 1996). Food products such as cake
and bread treated with >2% ethanol (wt/wt) were rejected by
consumers on the basis of flavour and/or odour (Seiler, 1978). A
level of 2% ethanol in food can extend the mould free shelf-life of
products but is generally not sufficient to prevent mould
development.
The inhibitory effect of ethanol can also be obtained by using an
encapsulated ethanol pouch (Ethicap, Freund Industrial Co. Ltd.,
Tokyo, Japan). This ethanol emitter extended the shelf-life of
packaged apple turnovers (Smith et al., 1987), pita bread (Black,
Quail, Reyes, Kuzyk, & Ruddick, 1993), packed sliced rye bread
(Salminen et al., 1996), English-style crumpets (Daifas et al., 2003)
and pre-baked buns (Franke, Wijma, & Bouma, 2002). The effects of
ethanol on a complex mycoflora were difficult to assess because the
ethanol in the headspace was not constant throughout the experiments. This was due to the absorption of ethanol by the products.
Therefore the effect of ethanol could also be attributed to the
ethanol absorbed by the product. It was shown that the concentration of ethanol required for inhibition was dependent on the
water activity of the system (Smith et al., 1987). In fact, it was
shown that both aw and moisture affected the vapour pressure of
ethanol as a consequence of water-solute and ethanolesolute
interactions in the matrix. These interactions varied according to
the modality of equilibration (desorption or absorption) at a given
aw (Pittia et al., 2006).
In addition, the use of ethanol emitter was related to modified
atmosphere packaging and it was suggested that ethanol vapours
could also affect the film’s permeability to oxygen (Black et al.,
1993). The empirical studies that have used ethanol for either
increasing the mould-free shelf-life of food products or controlling
postharvest decay of fruits are reported in Table 1.
362
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
Table 1
List of empirical studies that have used ethanol for either increasing the mould free
shelf-life of food products or controlling postharvest fruit decay.
Product
Treatment
Concentrations
Citation
Table grapes
Table grapes
Peaches
Pre-baked buns
Vapour
Vapour
Vapour
Ethanol
emitter
Incorporation
Immersion
Immersion
Immersion
Immersion
Immersion
Vapour
5 ml/kg grape
2 ml/kg grape
20e45%
0.55e2.2 g/140 g
bun
2e12% (v/v)
35, 50% (v/v)
20, 50%
10, 20%
20e70%
20e50%
20e70%
Chervin et al., 2003
Chervin et al., 2005
El-Sheik Aly et al., 2000
Franke et al., 2002
Geiges & Kuchen, 1981
Karabulut et al., 2003
Karabulut et al., 2004
Karabulut et al., 2005
Lichter et al., 2002
Lichter et al., 2005
Lihandra, 2007
Vapour
Immersion
Immersion
4, 8 ml/kg grape
5e20%
10, 20%
Lurie et al., 2006
Margosan et al., 1994
Margosan et al., 1997
35%
0.5e3.5%
10, 20%
0.33e1.65 g/215 g
bread
0.2e1.4%
0.5e3.5%
2.5e40%
2.2 g/100 g apple
Mlikota Gabler et al., 2005
Plemons et al., 1976
Romanazzi et al., 2007
Salminen et al., 1996
Seiler, 1984
Seiler & Russell, 1991
Smilanick et al., 1995
Smith et al., 1987
10, 20% (w/v)
0.5e2%
0.06e0.16% (v/v)
200e1500 mL/L
Spadaro et al., 2004
Vora & Sidhu, 1987
Yuen et al., 1995
Zhang et al., 2007
Bread
Grapes
Table grapes
Table grapes
Table grapes
Grapes
Oranges and
peaches
Table grapes
Strawberries
Peaches and
nectarines
Table grapes
Bread
Table grapes
Sliced rye bread
Immersion
Incorporation
Immersion
Ethanol
emitter
Bread
Incorporation
Bread
Incorporation
Lemons
Immersion
Packaged apple Ethanol
turnovers
emitter
Apples
Immersion
Bread
Incorporation
Oranges
Vapour
Chinese
Vapour
bayberries
solutions 5e10% at 22 C for 1e4 min had no effect on the decay due
to the prevalent Rhizopus stolonifer, the decay was reduced from
77% (control) to only 1% after treatment with 10% ethanol for 4 min
at 45 C, Table 2. But, the same treatment did reduce the decay to
only 36% for “Swede” strawberries infested by the prevalent
Botrytis cinerea (Margosan et al., 1994). These results clearly indicate that the sensitivity to ethanol is greatly dependent on the
mould.
In a more recent study, peaches dipped into 20%e100% ethanol
solutions were completely rotten by ten days when stored at room
temperature, but the peaches experienced little to no browning. In
contrast, untreated and fungicide-treated fruit were protected for
1d and 2d, respectively (Lihandra, 2007). The effects of ethanol
vapour on the growth of Penicillium italicum and Penicillium digitatum on oranges were assessed by Yuen, Paton, Hanawati, and
Shen (1995). These authors reported that exposure to 0.16%
ethanol vapour for 5d delayed the appearance of infection symptoms caused by P. digitatum and P. italicum by about 10d and 8d,
respectively.
The use of ethanol in food at low concentrations (approximately
2% wt/wt) may inhibit growth and germination thus extending the
free shelf-life of food products. In contrast, at higher ethanol
concentrations, it is suggested that a significant fraction of the
spores present at the surface of the dipped fruits were inactivated.
However the exposure to ethanol did not appear sufficient for
a complete inactivation because fruits were eventually rotten. The
more recent studies aimed at optimizing the combination of
ethanol concentration, temperature and duration of the treatment
without altering the quality of fruits.
3. Liquid treatments
3.1. Effect of ethanol on fungal growth
2.2. Control of postharvest decay of fruits
Ethanol dips and vapours were reported to control postharvest
diseases of apple (Spadaro, Garibaldi, & Gullino, 2004), peaches (ElSheik Aly, Baraka, & El-Sayed Abbass, 2000; Feliciano et al., 1992;
Margosan et al., 1997), lemons (Smilanick, Sorenson, & Henson,
1995), Chinese bayberries (Zhang et al., 2007) and table grapes
(Chervin et al., 2003; Chervin, Westercamp, & Monteils, 2005;
Karabulut et al., 2003; Karabulut, Romanazzi, Smilanick, & Lichter,
2005; Lichter et al., 2002; Lichter, Zutahy, Kaplunov, Aharoni, &
Lurie, 2005; Lurie et al., 2006; Mlikota Gabler & Smilanick, 2001;
Mlikota Gabler, Smilanick, Aiyabei, & Mansour, 2002; Mlikota
Gabler, Smilanick, Ghosoph, & Margosan, 2005; Romanazzi et al.,
2007) especially when heated (Margosan, Smilanick, & Simmons,
1994; Margosan et al., 1997; Smilanick et al., 1995). Whereas
immersion of naturally-infested “Chandler” strawberries in ethanol
Geiges and Kuchen (1981) have assessed the influence of ethanol
on mycelium development for some moulds grown in pure cultures.
Experiments were carried out by adding ethanol to bread slices
inoculated with spores. Growth inhibition was observed qualitatively for ethanol concentrations in the range 0e12% (v/v) depending
on the mould. In the range 0e4%, growth was observed for all the
moulds studied (i.e., Penicillium expansum, Penicillium implicatum,
Penicillium lanoso-coeruleum (¼ Penicillium aurantiogriseum),
Aspergillus oryzae and Trichoderma harzianum). But growth was
arrested at 8% for all these moulds. In contrast, Perry and Beale (1920)
found that Penicillium glaucum (¼ most probably P. expansum) would
grow in concentrations up to 8% alcohol in dextrose broth. Another
study reported that the inhibitory concentration of ethyl alcohol for
Penicillium carmine-violaceum (¼ P. roseopurpureum), P. citrinum and
P. glaucum (¼ most probably P. expansum) were in the range 5e5.5%
Table 2
Effect of hot ethanol treatments on postharvest control of gray mould (Botrytis cinerea) and black rot (Rhizopus stonolifer) of strawberries. Selected data from Margosan et al.
(1994).
Strawberry (Control test)
Prevalent pathogen
Test
T ( C)
Decay (%)
Immersion time (min)
Chandler (Untreated Dry Control: 77% decay)
Rhizopus stolonifer
Swede (Untreated Dry Control:97% decay)
Botrytis cinerea
Water
10% ethanol
Water
10% ethanol
Water
10% ethanol
Water
10% ethanol
22
22
45
45
22
22
45
45
1
4
77
72
55
29
99
100
97
96
78
80
10
1
100
99
62
36
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
(Krause & Ellis, 1937). Biomass production by Pyrenophora avenae
was reduced by 20% in 1% ethanol, growth in 3% and 6% ethanol
reduced biomass by 57 and 95% respectively (Walters, McPherson, &
Cowley, 1998).
A non-linear model was developed for assessing the effect of
ethanol on the growth of twelve different moulds on Potato
Dextrose Agar (0.99 aw) at 25 C (Dantigny, Guilmart, Radoi,
Bensoussan, & Zwietering, 2005). This model was able to estimate
the minimum inhibitory concentrations, MIC for growth and the
concentrations of ethanol at which the radial growth rate, m was
KðMICEÞ
; where mopt was the
equal to mopt/2, K : m ¼ mopt K:MIC2K:EþMIC:E
growth rate at 0% ethanol. The model fit the experimental data with
a good accuracy. It was capable of describing curves, m vs. E, with
either a concave shape (K < MIC/2) or a convex one (K > MIC/2).
MIC was estimated in the range 3%e5% for all moulds with the
notable exception of T. harzianum (MIC ¼ 2.14%) and Paecilomyces
variotii (MIC ¼ 6.43%), Table 3. These results demonstrated that
except T. harzianum, no moulds were completely inhibited by
ethanol concentrations of 2% in foods. Aspergillus niger and Mucor
racemosus were characterised by equivalent minimum inhibitory
concentrations of about 4.2%. But the K values were fairly different,
3.82% and 1.57% for A. niger and M. racemosus, respectively. While
2% ethanol would decrease the growth rate of M. racemosus to
about a third of mopt, this concentration would have almost no
inhibition effect on A. niger.
3.2. Effect of ethanol on germination
In studies assessing the effect of ethanol on germination, spores
are exposed to ethanol throughout the experiments. These studies
aimed at determining the minimum inhibitory concentration for
germination. The underlying concept is that prevention of germination would also prevent from growth. Because many studies
assessing the influence of ethanol on fungal growth were carried
out using spores as the inoculum, it can be assumed that no growth
was observed because spores were unable to germinate. Accordingly, no difference was shown between the minimum inhibitory
concentrations for growth and for germination for some Penicillia
(Krause & Ellis, 1937). A complete inhibition of germination of
conidia from these Penicillia was achieved at 50 g l1. However,
it would be possible that some spores (conidia) were germinated
but eventually were no longer capable of producing hyphae. While
the minimum ethanol concentration for growth of Penicillium
chrysogenum was estimated to 3.93% (Dantigny,Guilmart et al.
2005), the minimum ethanol concentration for germination was
estimated to 4.3% (Dantigny, Tchobanov, Bensoussan, & Zwietering,
2005). In the latter study, some spores were producing a germ tube
Table 3
Minimum inhibitory concentrations (MIC) and ethanol concentrations at which the
growth rate is halved, (K) for various food spoilage fungi. Adapted from Dantigny,
Guilmart, et al. (2005)
Mould
K (%, wt/wt)
MIC (%, wt/wt)
Aspergillus candidus
Aspergillus flavus
Aspergillus niger
Cladosporium cladosporioides
Eurotium herborarium
Mucor circinelloides
Mucor racemosus
Paecilomyces variotii
Penicillium chrysogenum
Penicillium digitatum
Rhizopus oryzae
Trichoderma harzanium
2.13
1.57
3.82
1.86
1.71
1.43
1.33
3.25
2.22
0.71
1.63
1.16
3.92
4.60
4.22
3.58
3.25
4.09
4.13
6.43
3.93
3.36
4.79
2.14
363
(synonymous for germination) that eventually blown up. More
interestingly, after 3 weeks, the cultures that failed to germinate
were allowed to continue to incubate. The ethanol solutions of
4e6% (wt/wt) were substituted for a 9 g/L NaCl solution, and the
cultures were incubated at 25 C. In all cases, visible mycelium
appeared during the next days, indicating that many conidia
remained viable. Under these experimental conditions, the inhibitory effect was clearly reversible.
3.3. Influence of ethanol on inactivation
Micro-organisms should be killed rather than inhibited, so it is
important to know which ethanol concentrations will result in
inactivation. It was shown that exposure to 40% ethanol completely
inhibited the germination of B. cinerea (Lichter et al., 2002), but the
immersion time was not stated. In another study, it was reported
that no germination of B. cinerea was observed on potato dextrose
agar (PDA) after immersion in 30% ethanol for 10s at 22e24 C
(Karabulut et al., 2004). Germination of B. cinerea spores on PDA
after a 30s immersion in 10 or 20% ethanol was 87 and 56%,
respectively, compared to 99% among untreated controls
(Karabulut et al., 2005). The incidence of grey mould (caused by B.
cinerea) infected grape berries that were untreated, or immersed
for 1 min in ethanol 35% at 25 or 50 C, was 78.7, 26.2, and 3.4
berries/kg, respectively, after 1 month of storage at 0.5 C and 2
days at 25 C (Mlikota Gabler et al., 2005).
Fifty percent ethanol can inactivate Sclerotina fructicola conidia
in 0.08 min on peach fruit surfaces and 60% ethanol killed Gilbertella persicaria and R. stolonifer sporangiospores on uninjured
surfaces of peach fruits in 1min (Ogawa & Lyda, 1960). But the same
authors reported that, on injured surfaces, 70% ethanol for 40 min
was needed to kill Gilbertella and 70% for 60 min to kill Rhizopus.
Margosan et al. (1994) demonstrated a synergistic effect of heat and
ethanol when they reported that diluted ethanol treatments
controlled postharvest decay of strawberries caused by B. cinerea
and R. stolonifer. A similar approach, employing exposure to
10e20% (v/v) ethanol at 46e50 C for up to 2.5 min, controlled
green mould on lemons, caused by P. digitatum and brown rot
caused by Monilinia fructicola on peaches and nectarines (Margosan
et al., 1997; Smilanick et al., 1995).
Temperature has marked influence on ethanol toxicity and its
effectiveness to control postharvest decay. Heat and ethanol
combinations were synergistic in the control of some fungi (Dao,
Bensoussan, Gervais, & Dantigny, 2008) and the diseases they
cause facilitating the use of lower temperatures and reduced
ethanol concentrations when these approaches are combined. The
influence of temperature during a 30 s exposure on the in vitro
toxicity of ethanol, expressed the concentrations (%, vol/vol) that
inhibited the germination of the treated population by 50% (EC 50)
was reported by Mlikota Gabler, Mansour, Smilanick, and Mackey
(2004). At 25 C, EC50 concentrations to R. stolonifer, A. niger, B.
cinerea and Alternaria alternata were 40, 36, 25, and 20% respectively, while at 40 C, the EC50 concentrations were 20, 20, 14, and
10% respectively. Ethanol toxicity to conidia of these fungi
increased about two-fold between 25 and 40 C.
By applying these treatments, injury to the fruits did not occur,
no off-flavours or odours were detected by the investigators, and an
increased firmness of the fruits was observed in most tests
(Margosan et al., 1997). These authors also reported that peaches
dipped in 20%e100% ethanol experienced little to no injury to the
fruit. Ethanol vapours improved apple appearance because they
were shown to reduce scald on apples (Scott, Yuen, & Ghahramani,
1995).
A complete inhibition of germination does not mean necessarily
that all spores are inactivated. Non-thermal inactivation, such as
364
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
that caused by ethanol, of a homogeneous population of microorganisms can be described with a D-value. The D-value, the
decimal reduction time, is the time that is needed to inactivate 90%
of the micro-organisms. From the data of Geiges and Kuchen
(1981), the D-values were calculated and reported in Table 4 for
different percentage of ethanol. In contrast to the other studied
mould, P. lanoso-coeruleum (¼ P. aurantiogriseum) was insensitive to
10% ethanol. Ethanol was effective from 13% and beyond. T. harzianum was the less resistant mould to ethanol in accordance with
previous results on the influence of ethanol on mould growth
(Dantigny,Guilmart et al. 2005). At 19% ethanol, P. implicatum was
the most resistant to ethanol, thus suggesting a greater z-value. The
z-value (% ethanol) can be considered as the increase in ethanol
concentration achieving the same inactivation within 10% of the
previous exposure time.
The water activity due to ethanol, awEtOH, and the water activity
due to glycerol, awGly, can be derived from the Langmuir equations
(Lerici, Nicoli, & Manzocco, 1996):
mH2 O
0:438awEtOH
¼
mEtOH
1 0:99awEtOH
(4)
0:236awGly
mH2 O
¼
mGly
1 0:99awGly
(5)
The water activity of the solution is given by the Ross equation:
awSol ¼ awEtOH $awGly
(6)
The relative humidity RH (%) in the headspace depends on the
water activity of the solution according to:
4. Vapour treatments
RH ¼ 100awSol
4.1. Thermodynamic relationship
Solving the equations, the composition of ternary solutions can
be calculated for setting the ethanol vapour pressure and the
relative humidity at the desired values physics permitted.
In a hermetically closed vessel, there are relationship between
the headspace and the liquid solution at steady state. Therefore it is
not necessary to measure the vapour pressure of ethanol in the
headspace. The liquid may be a binary water/ethanol or a ternary
water/ethanol/glycerol solution. In addition to modifying the
ethanol vapour pressure, the addition of glycerol allows the control
of the relative humidity in the headspace. To maintain the same
humidity in the headspace at different ethanol concentrations in
the liquid it is necessary to adjust the aw of the solution by adding
glycerol (or other aw depressant).
For a ternary solution that contains water (g), mH20; ethanol (g),
mEtOH; and glycerol (g), mGly, the molar fraction of ethanol in the
liquid is :
Xe ¼
mEtOH
46
(1)
mEtOH mGly mH2 O
þ
þ
46
92
18
The ethanol vapour pressure, Pe (kPa), in the headspace is:
Pe ¼ Xe ge Pt
(7)
4.2. Effect of ethanol on fungal growth
Exposing fruit to ethanol vapours proved effective at inhibiting
fungal growth. Peaches that were exposed to 70% or 100% ethanol
vapours were protected against fungal infection for up to 30 days
when stored at either 4 C or room temperature. In contrast, at
room temperature, untreated peaches and fungicide-treated
peaches were spoiled by fungi after 2 and 3 days, respectively. 20%
ethanol protected peaches for ten days when stored at 4 C and two
days at room temperature. Oranges that were exposed to 20%, 50%,
70% and 100% ethanol vapours were protected from fungal infection for 30 days at both 4 C and room temperature (Lihandra,
2007). The use of ethanol vapours at a dose rate of 2 ml kg 1 to
limit the development of B. cinerea on table grapes was also
reported (Chervin et al., 2005). The total inhibition of the growth of
Penicillium notatum, corresponding to the MIC of ethanol in vapour
phase, was observed at a concentration of 8.6 mmol/L air (Tunc,
Chollet, Chalier, Preziosi-Belloy, & Gontard, 2007).
(2)
where ge the activity coefficient that can be considered equal to 1
for Xe less than 0.8 (Clausen & Arlt, 2002), Pt (kPa) the ethanol
vapour pressure at saturation.
Pt depended on T ( K) according to a Claudius-Clapeyron law
that was approximated by plotting Ln (Pt) vs. (1/T), [data from Lide
(1995)]:
Ln ðPtÞ ¼ 19:11 5094=T
(3)
C,
At 25
(298 K) Pt ¼ 7.5 kPa. Assuming the atmospheric pressure 105 Pa, 1 kPa ethanol vapour pressure is equivalent to 1% (v/v)
ethanol in the headspace.
Table 4
Influence of ethanol on the inactivation of conidia of different fungi. D-values were
calculated from the data of Geiges & Kuchen, 1981.
Mould
D6% (d) D8% (d) D10% (d) D13% (d) D16% (d) D19% (d)
Penicillium expansum
Penicillium implicatum
Penicillium aurantiogriseum
Aspergillus oryzae
Trichoderma harzianum
ni
34.7
ni
ni
ni
ni
15.3
ni
87.2
16
11.8
11.1
ni
19.2
3.3
3.1
9.7
36.1
4.7
2.8
ni: less than 1log reduction in viable conidia after 100d.
1.7
7.8
13.1
2.7
0.9
0.8
3.9
1.4
1.4
0.5
4.3. Influence of ethanol on germination
The first detailed study of the inhibitory effects of ethanol
vapour on the germination of P. glaucum (¼ most probably
P. expansum) and Sterigmatocystis nigra was early demonstrated by
Lesage (1895, 1897). The inhibition of the germination of P. chrysogenum conidia by ethanol vapours was studied in the range 0e4%
(wt/wt) (Dantigny, Tchobanov et al. 2005). It was shown that all
conidia of P. chrysogenum germinated at ethanol concentrations
that were 3.0% (wt/wt) or less. At 3.5% (wt/wt), about 60% of the
inoculated spores were capable of germinating after 40 h, eventually forming a mycelium. After 3 weeks of incubation, no germination had occurred at ethanol concentrations of 4% and higher.
Ethanol vapour was also effective in retarding the apparition of
green mould and blue mould on oranges (Yuen, Paton, Hanawati &
Shen, 1995). The authors reported that exposure to 0.16% ethanol
vapour for 5d delayed the appearance of symptoms caused by P.
digitatum and P. italicum, by 10d and 8d respectively. It was suggested that growth was delayed due to an inhibition of the
germination.
The effectiveness of ethanol vapour for inhibiting growth and
germination of fungi is well demonstrated. However, all the
experiments were carried out on products and media that contained water. Therefore, ethanol in the headspace would have been
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
absorbed by these products and media. Mycelium growing at the
surface would use nutrients and ethanol present in the medium.
Therefore the real cause for inhibition could be the ethanol absorbed by the medium rather than ethanol vapour itself. The same
phenomenon would have occurred also during germination,
although to a lesser extent because interactions between the spore
and the medium at the early stage of germination are limited.
4.4. Influence of ethanol on inactivation of fungal spores
It was shown that at 23 C conidia of A. niger and P. notatum were
completely inactivated after being exposed to 25% ethanol for 3d
and 1d respectively (Bacílková, 2006). Similarly, more than 2 log
inactivation of conidia of some Penicillia was achieved by applying
20% (0.7 kPa) ethanol vapour in the headspace at 25 C for 1 d. An
increase of the inactivation up to 5 log was obtained for 40%, 1.5 kPa,
(Dantigny et al., 2007).
It was demonstrated that the effect of ethanol vapours on spore
inactivation depended on temperature. An increase in temperature
from 10 to 30 C was more important than an increase from 5 to 10%
(w/w) ethanol for explaining the inactivation of P. digitatum and P.
italicum (Dao, Bensoussan, Gervais & Dantigny, 2008). According to
the ClausiuseClapeyron law, an increase in temperature caused an
exponential increase in the ethanol vapour pressure. The increase
in the ethanol vapour pressure can be obtained also by decreasing
the water activity (Lerici et al., 1996). In fact, at reduced aw less free
water is available for binding ethanol resulting in an increase of
ethanol vapour pressure. Accordingly, the inactivation of spores of
P. chrysogenum, P. digitatum and P. italicum was greater at 0.70 aw
than at 0.90 aw (Dao et al., 2008). Therefore, the greater inactivation
of these spores at reduced water activity and high temperature
could be explained by an increase of the ethanol vapour pressure.
The relationship between the inactivation of enzymes by
ethanol and its vapour pressure was studied in a simple model
containing polyphenoloxidase, PPO, (Lerici & Manzocco, 2000).
Ethanol concentrations, water activity and temperature were
undoubtedly the dominant factors affecting efficiency of ethanol
towards PPO. It was demonstrated that the combined effects of
these variables were higher than the simple addition of the single
actions. This was attributed to an indirect effect of temperature
(and aw) on ethanol action and supported the hypothesis that
ethanol toxicity can be described by its vapour pressure.
The influence of the relative humidity (70e90%) and temperature (20e30 C) at a constant ethanol vapour pressure 0.6 kPa was
assessed for inactivating some Penicillia (Dantigny et al., 2007). At
70% RH, 30 C all conidia were viable after 24 h and 48 h exposure to
ethanol vapours, Table 5. Overall the greatest inactivation was
shown at 90% RH, 20 C. In this experimental condition the ethanol
concentration in the solution was the greatest, 23.40 g/100 g
solution. In contrast inactivation was not exhibited for the lowest
ethanol concentration, 7.99 g/100g solution. However, there was no
365
correlation between inactivation and the concentration of ethanol
in the solution. For all the species the inactivation was greater at
80% RH, 25 C (13.30 g ethanol/100g solution) than at 70% RH, 20 C
(15.22 g ethanol/100 g solution). In fact a more detailed analysis has
shown that RH was the key factor for explaining inactivation
(Dantigny et al., 2007). Temperature was also important, but to
a lesser extent.
The effect of ethanol vapours (range 0.3e0.45 kPa) on inactivation of conidia obtained by a standardised protocol that consisted
in preparing spore suspension was compared to that of dry-harvested conidia for some species of Penicillium. While all dry-harvested conidia remained viable after 24h treatment, about one
log10, 3.5 log10 and 2.5 log10 reductions were observed for hydrated
conidia of P. chrysogenum, P. digitatum and P. italicum respectively
(Dao & Dantigny, 2009). These results suggested that the intracellular water activity of the conidia may be correlated to the observed
differences in sensitivity to ethanol vapours. Ethanol is a very
hydrophilic molecule, and more ethanol could be dissolved in
a hydrated conidia during the exposure to vapours, eventually
inactivating the conidia. In contrast to conidia produced at the
laboratory, conidia found in the environment are usually not
hydrated. It was shown that at 2.8 kPa ethanol, more than 4 log10
reductions in viable dry-harvested conidia of P. chrysogenum,
P. digitatum and P. italicum were achieved after 24-h exposure (Dao,
Dejardin, Bensoussan, & Dantigny, 2010).
5. Mode of action of ethanol on fungi
The major target of ethanol as a stress agent is the cell membrane
of fungal cells. But it has many other effects, such as denaturation of
proteins (Mishra, 1993), and inhibition of the uptake of various
nutrients, e.g., the non-competitive inhibition of uptake of glucose
and ammonium ions (Leão & van Uden, 1982; Thomas & Rose, 1979).
The inhibition of fungal growth was partly attributed to the decrease
of aw due to ethanol. Water stress accounted for up to 31, 18 and 6% of
growth inhibition of A. oryzae by ethanol at 25, 40, and 42.5 C
respectively (Hallsworth, Nomura, & Iwahara, 1998).
The main function of the cell membrane of micro-organisms is to
form a permeability barrier, regulating the passage of solutes
between the cell and the external environment. In the environment,
micro-organisms may be confronted with lipophilic compounds
such as alkanols which preferentially accumulates in membranes.
This accumulation will affect physicochemical properties of
membranes and consequently their functioning (Weber & de Bont,
1996).
Lipophilic compounds, which possess a high affinity for the cell
membrane are more toxic than less lipophilic compounds. For
instance, ethanol is only toxic for micro-organisms at high concentrations (several %), whereas solvents like toluene are already toxic
in the mM range (Heipieper, Weber, Sikkema, Keweloh, & de Bont,
1994). However, this correlation does not apply for hydrophobic
Table 5
Influence of relative humidity and temperature on inactivation of conidia of some Penicillium species at a constant ethanol vapour pressure Pe ¼ 0.6 kPa. Adapted from Dantigny
et al. (2007).
RH (%)
T ( C)
EtOH (g)
H2O (g)
Gly (g)
log (N0/Nt) mean sd
P. chrysogenum
24H
70
90
80
70
90
20
20
25
30
30
15.22
23.40
13.30
7.99
12.24
41.18
76.40
53.44
38.31
71.63
43.60
0.20
33.26
53.70
16.12
0.00
3.96
3.72
0.00
3.68
P. digitatum
48H
0.00
0.04
0.05
0.00
0.05
2.33
4.26
3.76
0.00
3.80
24H
0.05
0.02
0.10
0.00
0.10
2.86
5.73
4.58
0.00
5.49
P. italicum
48H
0.58
0.46
0.17
0.00
0.24
2.91
6.04
5.96
0.00
6.43
24H
0.81
0.00
0.00
0.00
0.00
2.66
4.24
4.25
0.00
4.22
48H
0.03
0.06
0.04
0.00
0.09
2.89
4.77
4.74
0.00
5.45
0.19
0.08
0.10
0.00
0.04
366
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
solvents that are not generally toxic for micro-organisms (Inoue &
Horikoshi, 1991; Vermuë, Sikkema, Verheul, Baker, & Tramper,
1993). In fact, their poor solubility in water will prevent them from
reaching high concentrations in the membrane (Osborne, Leaver,
Turner, & Dunnill, 1990). The maximum concentration of a solvent
in the aqueous phase decreases and the hydrophobicity increases
with increasing the carbon chain length (Weber & de Bont, 1996).
Depending on the hydrophobicity of the solute, it will accumulate
more or less deeply into the bilayer. Alkanols like ethanol will
interact with the headgroup area and alkanes with the fatty acid
acyl-chains. The toxicity of alcohols is directly related to the length of
their aliphatic chain and their hydrophobicity (or lipophilicity). The
order of toxicity of alcohols to spores of Sclerotinia fructicola, R. stolonifer and G. persicaria was propanol, isopropanol, ethanol and
methanol (Ogawa & Lyda, 1960).
The addition of short-chain alcohols such as ethanol has a variety
of biophysical effects. Ethanol is a small molecule, which affects
the physical state and biological functions of cell membranes.
It interacts with membranes at the lipidewater interface, weakening the hydrophobic barrier to the free exchange of polar molecules, thereby perturbing membrane structure and function. This
could result in an increase in the surface area occupied by each
phospholipids molecule and a decrease in membrane thickness,
again increasing membrane permeability (Sikkema, de Bont, &
Poolman, 1995).
There is only one small proportion of ethanol that is distributed
in the membrane. When the ethanol is intercalated within the
hydrophobic part of the membrane, the polarity of this zone
increases and this authorizes the passage of other polar molecules
through the semi permeable membrane. Indeed, during insertion
in the double-layered one on the level of the polar heads, it has
been demonstrated that short-chain alkanols (C 3) can promote
the formation of an unusual phospholipid aggregation structure,
the interdigitated phase (Simon & McIntosh, 1984; Slater & Huang,
1988; Vierl, Löbbecke, Nagel, & Cevc, 1994). In the interdigitated
phase the lipid acyl-chains from the opposing monolayers are fully
interpenetrated, thereby exposing the terminal methyl groups.
These short-chain alkanols will anchor with their polar moiety to
the phopholipid headgroup, and with the non-polar part between
the phospholipid acyl-chains, Fig. 1. Since non-polar moiety of
these molecules is short compared to the fatty acid acyl-chains,
these molecules would potentially cause voids between the lipid
chains in the bilayer interior. As the energy of formation of holes
between hydrocarbons is extremely large, the lipids respond by
forming the interdigitated phase (Simon & McIntosh, 1984). As
polar headgroups are important for the barrier properties of the
membrane bilayer, it is expected that the formation of such lipid
phase will result in an increase permeability of the membrane.
Accordingly, a decrease in the osmotic water permeability of the
cell membrane has been observed with increasing concentration of
alcohol (25% decreases at 2% ethanol and 60% decrease at 5%
ethanol) with 1.0e1.25 M ethanol required to induce loss of semipermeability (Jones, 1989). Although the interdigitated lipid phase
Fig. 1. Schematic drawing, showing the effect of ethanol on the conversion of lipid
bilayers from the non-intergitated gel phase (Lb) to the fully interdigitated gel phase
(LbI). Adapted from Weber and de Bont (1996).
can be induced by various additives it should be noted that this
lipid aggregation structure is only observed for bilayers in the gelphase (Simon & McIntosh, 1984; Slater & Huang, 1988; Vierl et al.,
1994). In micro-organisms, however, most of the phospholipids
are generally in the liquid-crystalline phase. It is therefore unclear if
the interdigitated phase will occur or can be induced by solvents in
microorganisms.
The membrane is besides a barrier also a matrix for various
important enzymes. These include enzymes involved in solute
transport, and enzymes participating in the electron transport
chains. The results of Thomas and Rose (1979) indicate a relationship between inhibition of growth and nutrient transport by
alcohol and membrane lipid composition. The alcohols themselves
are highly permeable to the cell membrane, and it is proposed that
the increased resistance to water flow is due to the interaction of
alcohol with the membrane causing a decrease in the size of the
water pores.
The membrane of spore is protected by a wall made up of several
layers enable him to resist adverse conditions. The wall of the spore
of fungi consisted in a polymer association of polysaccharides and
chitin-glucanes. The envelope of the spore contains also simple
sugars (galactose, mannose), cellulose, proteins and lipids. During
germination of P. chrysogenum, it was observed that at 3.5% ethanol
some conidia had produced abnormal germinating tube that
eventually ruptured at the apex level while other conidia were
inhibited at the swelling stage (Dantigny, Tchobanov et al. 2005).
These observations support the hypothesis that ethanol is responsible for the leakage of solutes across the membrane and for cell
lysis following decreased peptidoglycan cross-linking in the
growing cell wall (Ingram & Buttke, 1984).
Relatively high concentrations of ethanol are required to kill
fungal spores compared with vegetative bacteria (Larson & Morton,
1991). Unbalanced cytoplasmic permeability and cytosol leakage
ultimately leading to disintegration of the cell, was reported to be
efficient in ethanol concentrations varying between 50 and 80%,
with a maximum at 70% (Hugo, 1971; Russell, Hugo, & Ayliffe, 1992).
But mature, swollen spores (due to hydration) are more easily killed
than dry and dormant ones, implying that any rational sanitizing
technique aiming at sporicidal action should be whenever possible
be applied to matures ones (Borick & Pepper, 1970; Tomazello &
Wiendl, 1995). Denaturation caused by alcohols can affect the
proteins of the cell wall, cytoplasmic membranes and proteins
contained in the cytoplasm (Bacílková, 2006). According to CabecaSilva, Madeira-Lopes, and van Uden (1992), the primary sites of
action of both heat and ethanol are mitochondrial membranes. Or
since low concentrations of ethanol can lower the temperature at
which phospholipids undergo a phase change, the increases in the
spore mortality and decay control following the addition of ethanol
may have resulted from a lowering of the phase-change temperature of mitochondrial membranes of the spores under these
conditions (Karabulut et al., 2004).
6. Conclusions
Ethanol has GRAS status in the USA, and it is an approved
substance for use as a disinfectant or sanitizer in organic crop
production by USDA National Organic Program (2001). Interest in
developing alternatives to fungicides is high because they are being
lost due to fungicide resistance, regulatory issues, and growers or
consumers who may prefer not to use them (Margosan et al., 1997).
In this review the effectiveness of ethanol for controlling fungi in
foods was demonstrated. Many studies have shown the efficacy of
ethanol treatments to reduce fruit decay. Ethanol residues after
treatment of peaches and nectarines were low after storage
(Mlikota Gabler et al., 2005; Margosan et al., 1997) and should pose
T. Dao, P. Dantigny / Food Control 22 (2011) 360e368
a minimal regulatory issue (Anonymous, 1993). Therefore, the use
of ethanol (either liquid or vapour) would have a great impact on
both the economy and food quality of fruits and vegetables. The use
of ethanol for preserving foods is dependent on regulations. At 2%
ethanol, ethanol is not effective at controlling mould growth, but
can be used as an additional barrier. It is also uncertain whether
consumers would accept an ethanol emitter in the packaging.
However, ethanol is used as a solvent for spraying aroma on cakes
prior to packaging in the baking industry.
Many fungicides are based on alcoholic molecules. Ethanol
solutions at difference percentages are used as a disinfectant for
personnel or surfaces. There are many indications that lower levels
of ethanol were required in the vapour phase as compared to direct
addition to a medium or food for complete inhibition (Lihandra,
2007; Smith et al., 1987). Ethanol is a very hydrophilic molecule.
It was shown that the effect of ethanol is greatly dependent on the
degree of hydration of the spore (Dantigny et al., 2007). Therefore,
this factor should be carefully controlled prior to comparing the
effect of ethanol solution and vapour.
In cases of large volumes to be treated, or equipment susceptible
to corrosion, the use of ethanol solutions is prohibited. Ethanol
vapours may be an interesting alternative to ethanol solutions. One
major concern with the use of ethanol vapour is its explosive
potential under high vapour pressure. The lower flammability limit
is 3.3 kPa (Anonymous, 1993). In this review, it was shown that 20%
ethanol at 25 C (0.7 kPa) could inactivate significantly fungal
spores. However, the air in manned workplaces cannot contain
ethanol at more than 0.1 kPa (Anonymous, 1993). Moreover,
ethanol vapour could be used for the protection of cereal grains
against the degradation and the production of mycotoxin by fungi
during storage. In contrast to ethanol solutions, ethanol vapour has
the advantage of reaching any remote places of the silo, especially
humid places where moulds were more likely to develop. This is
a promising use of ethanol vapour for preventing from the development of toxigenic moulds during storage of cereals and grains. At
present, fumigation with ethanol vapour for decontaminating
a cooler room in a bakery was proved effective but should be
optimised. These are applications that are currently being
examined.
Acknowledgements
Ailsa Hocking is gratefully acknowledged for helpful information on the taxonomy of Penicillium species.
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