Emerging Mycotoxins: Beyond Traditionally Determined Food Contaminants

https://mail.google.com/mail/u/1/?tab=Cm#inbox?projector=1

Comment; Fungal organisms like all types of life forms have the capacity to evolve, mutate, change to meet the demands placed upon them in order to survive. This paper reviews numerous new toxins that have been produced and are being changed, altered and produced that we need to be aware of as they are modifications of known toxins with undocumented (at the moment) effects on other organisms—such as us!

■ INTRODUCTION
Filamentous fungi show a remarkable potential to produce
secondary metabolites. Although the exact number of fungal
metabolites is unknown, a signiﬁcant fraction of the ∼170,000
currently known natural products1 are of fungal origin, and it
can be assumed that many more compounds are yet to be
discovered. Fungal metabolites include important pharmaceut-
icals such as penicillin or statins, potent poisons such as
aﬂatoxins or trichothecenes, and metabolites that are both toxic
and pharmaceutically useful, such as the ergot alkaloids.2 As
fungi can grow on basically any organic material, also food and
feed are regularly contaminated with fungal metabolites.
Poisonous fungal metabolitescommonly referred to as
mycotoxinsare of major importance to food and feed safety.
One of the best deﬁnitions of mycotoxins states that they “are
natural products produced by fungi that evoke a toxic response
when introduced in low concentration to higher vertebrates and
other animals by a natural route”.3 In addition to animal
toxicity, mycotoxins may exert phytotoxic or antimicrobial
eﬀects. A variety of fungi are known to produce mycotoxins,
including Aspergillus, Fusarium, and Penicillium species.4 The
most important classes of mycotoxins5 include the highly
carcinogenic aﬂatoxins (e.g., aﬂatoxin B1, AfB1),6 trichothe-
cenes (e.g., deoxynivalenol, DON),7 fumonisins (e.g., fumoni-
sin B1, FB1),8 ochratoxin A (OTA),9 and zearalenone (ZEN).10
These toxins and several others are regulated in many countries
of the world11 after thorough risk assessment, taking into
account toxicity, occurrence, and consumption data as well as
economic and political considerations.
Sound analytical methods are key to ensuring food safety,
and dozens of new methods are developed and published each
year.12,13 Of those, liquid chromatography−mass spectrometry
(LC-MS) based methods are becoming more and more

popular, as they allow the sensitive simultaneous determination
of multiple fungal metabolites in many matrices. These
methods also sometimes reveal “surprising” ﬁndings, such as
known mycotoxins in untypical matrices (e.g., fumonisin B2 in
grapes14) or in unusual geographical regions (e.g., aﬂatoxins in
Europe15), for which global warming might partly be
responsible.16 LC-MS is of tremendous help in the discovery
of new mycotoxins (e.g., NX-toxins17), as well as masked18 or
other modiﬁed19 forms of mycotoxins. Moreover, “emerging
mycotoxins”, which are in the scope of this review, can occur in
high frequency and sometimes also in high concentrations in
cereals and in other food- and feedstuﬀs.
The term “emerging mycotoxins”, although often used
nowadays for certain fungal metabolites, is not clearly deﬁned.
One of the ﬁrst papers to use this term was published in 2008
and deals with the Fusarium metabolites fusaproliferin (FP),
beauvericin (BEA), enniatins (ENNs), and moniliformin
(MON),20 and the term has mostly been used for these
compounds ever since. However, in a more recent paper,
emerging mycotoxins were deﬁned as “mycotoxins, which are
neither routinely determined, nor legislatively regulated;
however, the evidence of their incidence is rapidly increasing”.21
According to this deﬁnition, many more fungal metabolites
with known (or at least suspected) toxicity would fall in the
category of emerging mycotoxins.

Special Issue: Public Health Perspectives of Mycotoxins in Food
Received: July 30, 2016
Revised: August 30, 2016
Accepted: September 6, 2016
Published: September 7, 2016

Review
pubs.acs.org/JAFC

This is an open access article published under an ACS AuthorChoice License, which permits
copying and redistribution of the article or any adaptations for non-commercial purposes.

Downloaded via 192.155.76.162 on November 14, 2018 at 21:36:52 (UTC).
See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

The aim of this review is to critically assess the toxicity and
occurrence of diﬀerent fungal metabolites that are currently not
regulated in food or feed. We deliberately chose the most
“interesting” fungal compounds based on the frequency of
occurrence and concentrations in food and feed, as well as
current scientiﬁc interest. The following sections will deal with
the Fusarium metabolites ENNs, BEA, MON, FP, fusaric acid
(FA), culmorin (CUL), and butenolide (BUT), the Aspergillus
metabolites sterigmatocystin (STE) and emodin (EMO), the
Penicillium metabolite mycophenolic acid (MPA), and the
Alternaria metabolites alternariol (AOH), alternariol mono-
methyl ether (AME), and tenuazonic acid (TeA).
■ FORMATION, TOXICITY, AND OCCURRENCE OF
EMERGING MYCOTOXINS
Enniatins. ENNs are cyclic hexadepsipeptides that are
produced by several Fusarium species, such as F. avenaceum,
F. oxysporum, F. poae, or F. tricinctum, which grow mainly on
cereals. In total, 29 species of ENNs are known.22 ENNs A, A1,
B (Figure 1), and B1 are most frequently detected in food and

feed. The toxicity of ENNs is based on their ionophoric
properties. They facilitate the transport of mono- or divalent
cations such as K+ or Ca2+ across membranes, thereby
disrupting normal physiological concentrations of these
ions.23 Low micromolar concentrations of ENNs were shown
to be cytotoxic to diﬀerent cell lines24−28 and to reduce the
motility of boar spermatozoa.29 Cell death was shown to be
mediated by the induction of apoptosis28,30,31 via the
mitochondrial pathway24,32 or by the induction of necrosis
linked to lysosomal damage.33,34 There are conﬂicting data on
whether35 or not31 ENNs trigger the production of reactive
oxygen species (ROS). Several studies found no genotoxic
eﬀect of ENNs.30,31,34 Interestingly, ENN B1 and T-2 toxin
showed an antagonistic toxic eﬀect on cultured intestinal
epithelial cells and intestinal explants, which indicates that ENN
B1 down-modulates the gastrointestinal toxicity of T-2 toxin.36
ENNs were shown to interact with proteins. In rat liver
microsomes ENNs inhibited the activity of acyl-CoA:ch-
olesterol acyltransferase.37 ENN B was shown to bind
calmodulin and to inhibit 3′,5′-cyclo-nucleotide phosphodies-
terase.38 ENNs furthermore interact with multidrug resistance
ATP-binding cassette (ABC) transporters. The overexpression
of P-glycoprotein (ABCB1) and breast cancer resistance
protein (ABCG2) counteracted the cytotoxic eﬀect of ENNs
on human cancer cell lines.39 Ivanova and co-workers40 found
that two multidrug resistance ABC transporters facilitate the
transport of ENN B1 across a monolayer of the human
intestinal cell line CaCo-2 in the apical to basolateral direction.
Furthermore, exposure to ENNs counteracted the protective
eﬀect of ABC transporters against cytotoxic compounds in

yeast41,42 and human cancer cell lines,39 and it may
consequently alter the bioavailability of pharmaceuticals in vivo.
Several studies investigated the in vivo toxicity of ENNs in
rodents. Most of these studies show low toxicity. A single dose
of 50 mg ENNs/kg body weight (BW) did not induce signs of
toxicity in rats.43 Oral administration of ENN A to mice at a
dosage of 1 mg/kg BW/day for 6 days or subcutaneous (sc)
administration of ENN A to mice at a dosage of 0.5 mg/kg
BW/day for 6 days did not elicit a toxic eﬀect.44 A daily
intraperitoneal (ip) ENN B dose of 5 mg/kg BW administered
for 9 days did not aﬀect BW, food intake, or behavior in mice.45
In rats fed an ENN A contaminated diet (465 mg/kg feed,
corresponding to 20.9 mg/kg BW/day) for 28 days, no eﬀects
on body and organ weight, histology of duodenum tissue,
biochemical blood parameters,46 or any visible signs of illness47
were found. Although altered peripheral blood lymphocyte
levels were detected, their consequences are yet unclear. In the
sole study that showed toxic eﬀect of ENNs, ip administration
of 10−40 mg/kg BW at an interval of 8 h was lethal for
immune-deﬁcient mice within 2−5 days, whereas lower levels
caused only weight loss.48
Two studies investigated the distribution of ENNs in
diﬀerent tissues of rats, and the highest levels were detected
in jejunum, liver, and fat tissue.45,46 These ﬁndings led the
authors to suggest that the jejunum is a site of ENN
absorption46 and that ENNs primarily accumulate in fat-rich
tissues owing to their lipophilic properties.45 Dietary ENNs
were also shown to accumulate in the meat, skin, and liver of
broilers,49 and traces of the toxin were detected in a high
fraction of Finish egg samples.50 ENNs were shown to cross the
blood−brain barrier in mice.51
Bioavailability and toxicokinetics of ENN B were investigated
in pigs.52 The toxin had a high (∼91%) oral bioavailability and
was rapidly absorbed, distributed, and eliminated. Hence, the
authors suggested that rapid metabolization rather than low
bioavailability may explain the low in vivo toxicity of ENNs.
Several metabolites of ENNs have been reported. Mono-
oxygenated, dioxygenated, and N-demethylated metabolites
were detected in liver microsomes of rats, dogs, humans, and
chickens.53−55 The same metabolites were detected in liver,
plasma, and egg samples of chickens54 and in liver and colon
samples of mice.45
A mixture of enniatins termed “fusafungine” is used as a
nasal/oromucosal spray with antibiotic and anti-inﬂammatory
properties.56,57 However, in April 2016, the European
Medicines Agency of the European Union recommended to
revoke the authorization of fusafungine sprays “due to serious
allergic reactions and limited evidence of beneﬁt”.58
ENNs were detected in 37, 68, and 76% of food (n = 4251),
feed (n = 3640), and unprocessed grain (n = 2647) samples
collected in Europe between 2000 and 2013.59 Maximum
reported concentrations in grains were 950, 2000, 18,300, and
5720 μg/kg for ENN A, ENN A1, ENN B, and ENN B1,
respectively. Maximum reported concentrations in cereal-based
food were, 42, 125, 832, and 980 μg/kg for ENN A, ENN A1,
ENN B, and ENN B1, respectively. ENNs were detected in
96% of samples (n = 83) of feed and feed raw materials with
median and maximum concentrations of 30 and 5441 μg/kg,
respectively.60 In a survey of Chinese medicinal herbs, ENN A,
ENN A1, ENN B, and ENN B1 were detected in 8, 7, 12, and
7% of all analyzed samples (n = 60) with maximum
concentrations of 355, 253, 291, and 40 μg/kg, respectively.61

Figure 1. Chemical structure of enniatin B.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7053

In summary, ENNs are toxic in vitro, but most in vivo data
indicate no or only low toxicity. ENNs are frequently detected
in food, feed, and grains, but feeding studies conducted with
rodents indicated a high tolerance to dietary ENN levels 1
order of magnitude higher than the reported maximum levels.
In their scientiﬁc opinion from 2014, the European Food Safety
Authority (EFSA) concluded that acute exposure to ENNs is
not a concern to human health, whereas no conclusion could be
drawn with respect to chronic exposure due to the lack of
relevant in vivo toxicity data.59 Interestingly, eﬀects of ENNs on
the bioavailability of pharmaceuticals have been suggested by in
vitro data and should be further clariﬁed before any valid
conclusions on that aspect can be drawn.
Beauvericin. BEA is a cyclic hexadepsipeptide (Figure 2),
produced by several Fusarium species, such as F. proliferatum,

F. subglutinans, F. verticillioides, or F. oxysporum. BEA is an
ionophore; it transports mono- or divalent cations, for example,
K+ or Ca2+, across membranes and thereby disrupts normal
physiological concentrations of these ions.62,63 Accordingly, low
micromolar concentrations of BEA were shown to be cytotoxic
to diﬀerent cell lines in vitro.25,26,31,64 Cell death was shown to
be mediated by apoptosis31,65−67 via the mitochondrial
pathway32,68 or by necrosis.69 There are conﬂicting reports
on whether35,70 or not31 oxidative stress is involved in BEA
toxicity. BEA showed no mutagenicity in the Ames test71 and
intercalated into DNA only when applied at high concen-
trations (>100 μM).31 However, it showed a mutagenic
potential in the alkaline Comet assay.72 Low micromolar BEA
concentrations aﬀected the development of immune cells in
vitro. BEA disturbed the maturation process of dentritic cells
and decreased the endocytosis ability of macrophages when
applied during their diﬀerentiation process.73 The toxin was
furthermore shown to act as an enzyme inhibitor in liver
microsomes. It inhibited cytochrome P450 enzymes74 and acyl-
CoA:cholesterol acyltransferase, the enzyme that causes the
accumulation of cholesteryl ester in atherogenesis.37 BEA was
shown to interact with P-glycoprotein (ABCB1) and breast
cancer resistance protein (ABCG2) in vitro.39 The data suggest
that BEA is both a substrate and an inhibitor of the investigated
transporters. It was concluded that the presence of homologues
of these transporters in the gastrointestinal tract may decrease
the bioavailability of BEA. Furthermore, the inhibitory eﬀect of
BEA on these transporters may aﬀect the bioavailability of
pharmaceuticals.
In vivo, toxicity and pharmacological behavior of BEA were
studied in mice and poultry. A daily ip administration of 5 mg/
kg BW for 9 days did not aﬀect BW, food intake, or behavior in

mice.45 The LD50 values in mice were ≥100 and ≥10 mg/kg for
oral and ip administration, respectively.75 When the distribution
of BEA in diﬀerent tissues was investigated, the highest
concentrations were detected in the liver and in fat tissue,
suggesting an accumulation in fat-rich tissue due to the
compound’s lipophilic properties.45 BEA was shown to cross
the blood−brain barrier in mice.51 The eﬀect of chronic dietary
exposure to BEA was investigated in poultry. In turkey and
broiler feeding trials, diets contaminated with ≤2.5 and ≤12
mg/kg BEA, respectively, and cocontaminated with other
Fusarium mycotoxins, had no eﬀect on performance parame-
ters, biochemical blood parameters, or meat quality.76−79
However, increased heart weight was detected in one broiler
trial.76 Dietary exposure of broilers and laying hens to 10 and 9
mg/kg BEA, respectively, did not aﬀect growth, feed uptake, or
egg production.49 Surveys of poultry products in Finland found
trace amounts of BEA in a high fraction of analyzed egg
samples and in a low fraction of analyzed meat and liver
samples.50,80 The authors suggested that BEA accumulates in
egg yolk owing to its lipophilicity.
BEA was detected in 20, 21, and 54% of food (n = 732), feed
(n = 861), and unprocessed grain (n = 554) samples collected
in Europe between 2000 and 2013.59 Maximum reported
concentrations for BEA in grains and in cereal-based food were
6400 and 844 μg/kg, respectively. BEA was detected in 98% of
samples (n = 83) of feed and feed raw materials with median
and maximum concentrations of 6.7 and 2330 μg/kg,
respectively.60 In a survey of Chinese medicinal herbs, BEA
was detected in 20% of all analyzed samples (n = 60) with a
maximum concentration of 125 μg/kg.61
In summary, BEA isalthough toxic in vitronot toxic to
rodents and poultry in vivo. Dietary BEA concentrations
applied in poultry feeding trials were comparable to maximum
concentrations reported in feed and feed raw materials. In vivo
toxicity data for other species are missing. BEA presumably
accumulates in fat-rich tissue including eggs of laying hens.
However, detected levels should be negligible concerning an
acute risk to human consumers. In their scientiﬁc opinion from
2014, EFSA concluded that acute exposure to BEA is not a
concern to human health, whereas no conclusion could be
drawn with respect to chronic exposure due to the lack of
relevant in vivo toxicity data.59 Eﬀects of BEA on components
of the immune system and on the bioavailability of
pharmaceuticals were suggested by in vitro studies and should
be investigated in vivo.
Moniliformin. MON (1-hydroxycyclobut-1-ene-3,4-dione;
semisquaric acid; Figure 3) is a small, water-soluble molecule

originally isolated from a Fusarium strain, initially called F.
moniliforme, in the 1970s.81 MON is among the most acidic
organic acids and therefore occurs in nature typically as sodium
or potassium salt. It is produced by a large variety of Fusarium
spp. (e.g., F. proliferatum,82 F. subglutinans,83 F. avenaceum,82
F. tricinctum,84 F. fujikuroi,85 F. nygamai,86 F. pseudonygamai,87
F. temperatum,88 or F. thapsinum87) and only recently has been
shown to be a metabolite of Penicillium melanoconidium.89

Figure 2. Chemical structure of beauvericin.

Figure 3. Chemical structure of moniliformin.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7054

The primary mode of action of MON seems to be the
inhibition of thiamin pyrophosphatase dependent enzymes,
which compromises the tricarboxylic acid cycle. Soon after the
discovery of impaired oxidation of pyruvate and α-ketoglutarate
by MON,90 inhibition of pyruvate dehydrogenase was
demonstrated in vitro.91,92 Later, several enzymes sharing
thiamin as common cofactor (pyruvate dehydrogenase, α-
ketoglutarate dehydrogenase, pyruvate decarboxylase, and
acetohydroxy acid synthase) have been shown to be inhibited
by MON.93 The authors concluded that MON meets the
criteria for an active site-directed, irreversible aﬃnity label for
enzymes utilizing thiamin and that the inhibition of pyruvate
dehydrogenase is cofactor-directed and likely caused by the
structural similarity of MON and pyruvate.
MON was shown to by phytotoxic to wheat and corn, where
it caused necrosis, aﬀected growth regulation, and rendered
leaves distorted.81 Its toxicity on animal cells was tested
extensively in vitro, and its eﬀects are highly dependent on the
used cell lines. For instance, MON showed no inhibitory eﬀects
on the proliferation of human white blood cell progenitors or
human platelet progenitors, but was cytotoxic to human red
blood cell progenitors with a half-maximal inhibitory
concentration (IC50) value of 4.1 μM.94 After treatment of
human hepatocellular carcinoma cells for 2 or 3 days with
MON, IC50 values of 39.5 μg/mL (403 μM) and 26.8 μg/mL
(273 μM) were recorded.95 With regard to chicken primary cell
cultures, MON was not toxic to chondrocytes and macro-
phages, but was toxic to splenocytes, cardiac (IC50 = 95 μM),
and skeletal myocytes (IC50 = 42 μM).96 Literature reports on
genotoxicity are slightly ambiguous. MON did not show any
activity in gene mutation assays with Salmonella typhimurium or
Escherichia coli with or without metabolic activation.97
However, the same authors elucidated that MON caused
pronounced dose-dependent chromosomal aberration in
primary rat hepatocytes. Compared to the control, a 9-fold
increase was seen after 3 h of exposure to 1 μg/mL (10 μM).
Previously, studies with 5−500 μM MON on the same cell type
showed no genotoxic eﬀects as measured with the unscheduled
DNA synthesis test.98 The most recent study that investigated
genotoxic eﬀects of MON used human peripheral blood
lymphocytes.99 Chromosomal aberrations, sister-chromatid
exchanges, and micronucleus frequencies were signiﬁcantly
increased in a dose-dependent manner compared with the
negative control after treatment of lymphocytes with MON
concentrations between 2.5 and 25 μM.
MON shows even more severe eﬀects in vivo. Incomplete
uptake of the polar toxin through the cell membranes in cell
culture studies may be a possible explanation for this
discrepancy. LD50 values of MON have been reported as 4.0
mg/kg BW in 1-day-old cockerels (oral),81 as 2.2−2.8 mg/kg
BW in 9-month-old female mink (ip),100 as 1.4 mg/kg BW in 7-
week-old female broiler chickens intravenous (iv),101 as 20.9
and 29.1 mg/kg BW in female and male mice (ip),102 as 3.7
mg/kg BW in 7-day-old ducklings (oral),103 and as 41.6 and
50.0 mg/kg BW in female and male rats (oral).103 Besides
damage to the heart muscle, including myocardial lesions and
increased relative heart weights,100,103−105 also muscular
weakness,103,104 respiratory distress,103,104 decreased feed intake
and BW gains,101,105 and impaired immune function104,106 were
noted frequently. In summary, these studies highlight that
poultry are the most aﬀected species and that the heart is the
main target organ in all investigated animals. Potential
synergistic eﬀects of the cardiotoxic MON with the co-

occurring BEA and ENNs107 or FB1105,106 were disproved for
the investigated conditions.
Occurrence of MON in cereals seems to be rather frequent
in various regions of the world, albeit at rather low levels on
average. For instance, 40% of 151 Finnish cereal samples
collected in 2005 were positive for MON, with a mean level of
190 μg/kg and a maximum level of 850 μg/kg.108 The toxin
was also found in maize samples from all agroecological zones
of Lesotho from the 2009/2010 and 2010/2011 seasons with
levels reaching 1.2 mg/kg.109 Furthermore, 93% of naturally
contaminated maize samples from northwestern Italy (n = 108)
collected in 2008, 2009, 2010, and 2011 showed detectable
levels of MON with maximum concentrations of 2.6, 0.5, 0.9,
and 0.4 mg/kg for the respective years.110 MON was detected
in 26 of 50 samples of poultry feed mixtures of Slovak origin in
concentrations up to 1.2 mg/kg.111
Two excellent reviews compiled general information on
MON112 or data on the negative health eﬀects caused by
MON.113 The latter paper stated that “MON per se does not
pose a clear threat to human health at current levels”, if an
internal no observed adverse eﬀect level (NOAEL) of 10 mg/
kg BW/day was applied. However, a more recently published
study indicated a severe impact of MON on the immune
system of Sprague−Dawley rats and a lowest observed adverse
eﬀect level (LOAEL) value of 3 mg/kg BW.114 Also, it was
assumed a few years ago that MON “is not likely to be a
genotoxic carcinogen”,113 which at least has to be questioned
given the latest data,99 and further studies should be conducted
to answer this question. The European Commission Direc-
torate-General for Health and Consumers requested a scientiﬁc
opinion from EFSA about the risks for public health related to
MON in feed and food,115 which is due soon. Disregarding
potential genotoxicity, current average contamination levels of
cereals imply that the compound is likely not a problem for
human health. Still, guidance levels for MON in poultry feed
might be warranted to better protect animals from occasional
high concentrations and to provide scientiﬁc support to poultry
farmers.
Fusaproliferin. FP is a bicyclic sesterterpene (Figure 4). It
is produced by Fusarium species such as F. proliferatum,

F. subglutinans, and F. verticillioides. FP was moderately
cytotoxic to lepidopteran SF-9 cells and to human B
lymphocyte IARC/LCL 171 cells. In the case of SF-9 cells,
the CC50 values (concentration resulting in 50% cell viability)
were 100 and 70 μM after 24 and 48 h of exposure,
respectively.116 Fornelli and co-workers found a CC50 value
of >100 μM after 48 h of exposure.25 For IARC/LCL 171 cells
the CC50 values were 60−65 and 55 μM after exposure times of
24 and 48 h, respectively.116 Moreover, exposure to 30 μM FP
inhibited the proliferation of IARC/LCL 171 cells.116 When the
transport of FP across a monolayer of Caco-2 cells was
investigated, 80−83% of FP that had been applied to the apical
side was detected on the basolateral side after 4 h of incubation,
which suggests a high bioavailability in vivo.117

Figure 4. Chemical structure of fusaproliferin.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7055

In vivo toxicity data are limited to brine shrimp (Artemia
salina) larvae and chicken embryos. FP was toxic in the brine
shrimp larvae bioassay with an LD50 of 53.4 μM or 23.7 μg/
mL.116 FP exerted teratogenic and pathogenic eﬀects on
chicken embryos when applied at concentrations of 1 or 5
mM.118 The toxin was shown to noncovalently interact with
oligonucleotides in vitro.119 Although interactions with DNA
may account for the teratogenic eﬀect of FP, they have not yet
been conﬁrmed in living organisms or cells.
FP was detected in 17% of samples (n = 83) of feed and feed
raw materials with median and maximum concentrations of 2.6
and 14.8 mg/kg, respectively.60 A review of FP occurrence data
published between 1997 and 2011 shows its occasional
occurrence in grains and grain-based foodstuﬀs120 at concen-
trations up to 500 mg/kg.121
In summary, high levels of FP occasionally occur in grains
and grain-based food and feed. However, its toxicity and mode
of action have not been comprehensively investigated so far.
Toxic eﬀects on chicken embryos and brine shrimp larvae
suggest a potential hazard to humans and animals that should
be clariﬁed by further experiments.
Fusaric Acid. FA (5-butylpiconilic acid; Figure 5), is a
phytotoxin produced by a great variety of Fusarium species,

such as F. oxysporum, F. moniliforme, F. proliferatum, F. sub-
glutinans, F. verticillioides, F. crookwellense, F. napiforme, and
F. fujikuroi.122,123 This oﬀ-white to yellowish crystalline
compound, a precursor of the beta blocker bupicomide,124 is
understudied for its mode of action and its eﬀects in livestock
and humans. FA has been shown to exhibit cytotoxic and
growth inhibitory eﬀects in diﬀerent normal and cancer cell
lines. Cell proliferation was suppressed by 500 μM FA in
human WI-38 ﬁbroblasts and colorectal adenocarcinoma cell
lines.125 Furthermore, FA was cytotoxic to dog kidney (IC50 =
10 μg/mL), McCoy mouse (IC50 = 25 μg/mL), and Chinese
hamster ovary ﬁbroblast cells (IC50 = 10 μg/mL), as well as to
rat hepatoma cells (IC50 = 50 μg/mL).126 FA was shown to
induce oxidative and mitochondrial stress, as well as to cause
apoptosis and necrosis in the human hepatocellular carcinoma
cell line HepG2.127 May and co-workers found that FA inhibits
the growth of the rumen microorganisms Ruminococcus albus
and Methanobrevibacter ruminantium at concentrations >84
μM.128 Another in vitro study demonstrated that FA inactivates
several clinical and soil isolates of Acanthamoeba, with an IC50
value of 0.31 μM.129 As a phytotoxin, FA is known to play a
critical role in accelerating the development of Fusarium wilt in
numerous plants, such as banana and tomato plants.130−132
The LD50 values of FA in mice were 100 and 80 mg/kg BW
for iv and ip administration, respectively.133 The average oral
bioavailability of FA (25 mg/kg) in Sprague−Dawley rats was
58%.134 Several studies indicated a neurochemical eﬀect of FA.
Researchers demonstrated a hypotensive eﬀect of FA in
humans, rabbits, rats, cats, and dogs that was based on the
inhibition of dopamine-β-hydroxylase,133,135,136 the enzyme
that converts dopamine to norepinephrine. FA was furthermore
shown to alter brain and pineal neurotransmitters in rats.137

Oral administration of FA (200 mg/kg BW) to swine caused
neurochemical changes and vomiting behavior.138 Co-admin-
istration of diﬀerent doses of FA and DON to swine decreased
weight gain and feed consumption, which led the authors to
suggest that FA might act synergistically with trichothecenes.139
Chickens that received FA-contaminated diet (up to 150 mg
FA/kg diet), on the other hand, did not show any abnormalities
in behavior, feed intake, weight gain, and appearance of visceral
organs.140 FA exerted a teratogenic eﬀect in zebraﬁsh. The
compound caused malformations at the notochord due to
copper chelating.141 A synergistic cytotoxic eﬀect was detected
for in ovo administration of FA and FB1.142 Interestingly, oral
administration of 1 mg/mL FA inhibits tumor growth of head
and neck cancer in an in vivo murine model.143
Porter and co-workers showed that FA is a naturally
occurring contaminant in several types of cereal grain and
mixed livestock and poultry feeds and is synthesized together
with other Fusarium mycotoxins.137 Smith and Sousadias
reported that in 48 investigated swine feed samples, 85%
were positive, with the highest concentration of 136 mg/kg
FA.144
Because FA is produced by the most common fungal genus,
Fusarium, due to its neurochemical eﬀects and due to possible
synergistic eﬀects with predominant mycotoxins (DON and
FB1), this substance might pose a problem to humans and farm
animals. FA is not yet regulated in any country or monitored
regularly. For an appropriate risk assessment, more studies
covering in vivo and in vitro trials as well as occurrence are duly
needed.
Culmorin. CUL is a sesquiterpene diol (Figure 6) produced
by several Fusarium species, such as F. culmorum, F. graminea-

rum, F. crookwellense, and F. venenatum (summarized by
Pedersen and Miller145) and by the marine ascomycete
Leptosphaeria oraemaris.146 In addition, the formation of various
related compounds, namely, hydroxyculmorins (5-, 12-, 15-
hydroxyculmorin), culmorone, and hydroxyculmorones, has
been reported for diﬀerent Fusarium strains.147,148
Information on the toxicological relevance of CUL and its
hydroxylated forms is limited. CUL was shown to be
phytotoxic149,150 and to possess antifungal activity.146 Pedersen
and Miller demonstrated that this compound results negative in
the Ames test.145 In baby hamster kidney cells BHK-21, only
the highest tested CUL concentration (20 μg/mL; 84 μM)
exhibited mild cytotoxicity.147 The LD50 for CUL in the chick
embryotoxicity screening test (CHEST) was between 68.0 μg/
kg (incubation period of 22 days) and 78.2 μg/egg (incubation
period of 7 days).151 Thus, CUL was approximately 10 times
less toxic than DON and approximately 25−30 times less toxic
than STE in the same test. In an attempt to extrapolate data
from the CHEST, the mouse LD50 (ip) for CUL was estimated
to range between 250 and 1000 mg/kg BW, indicating low
mammalian toxicity.145
So far, in vivo studies on the toxicity of CUL are restricted to
caterpillars and swine. Administration of a CUL-contaminated

Figure 5. Chemical structure of fusaric acid.

Figure 6. Chemical structure of culmorin.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7056

diet (25 mg/kg) for 7 days did not aﬀect the mortality or
weight of corn earworm (Heliothis zea) and fall armyworm
(Spodoptera frugiperda) larvae.152 Similarly, CUL exposure (2
mg/kg diet, 21 days) had no negative impact on the
performance of growing piglets.153 Interestingly, CUL even
seemed to slightly enhance the growth of H. zea larvae and to
increase the weight gain of pigs. In addition, a potential
interaction of CUL and DON was investigated in both studies.
The combination of CUL (10 mg/kg diet) with DON (25 mg/
kg diet) resulted in a signiﬁcantly increased mortality and
decreased weight of H. zea larvae when compared to DON
alone. The authors assumed a synergistic eﬀect, which is
probably related to an impaired DON detoxiﬁcation in insects
in the presence of CUL.152 While an interaction of these two
mycotoxins could not be conﬁrmed in pigs,153 CUL was
recently proposed to increase DON toxicity in wheat seeds.150
The latter is of certain interest, because the levels of CUL as
well as hydroxyculmorins strongly correlate with those of DON
in cereal grains.154,155 In a recent study, CUL was detected in
95−100% of Norwegian barley, oat, and wheat samples (n =
76). Highest contamination levels were found in oats (n = 28),
where median and maximum CUL concentrations reached 2.0
and 31.5 mg/kg, respectively.155 Streit and co-workers found
63, 63, 13, and 7% of feed and feed raw materials (n = 83)
positive for CUL, 15-hydroxyculmorin, 5-hydroxyculmorin, and
15-hydroxyculmorone.60 In contrast, hydroxyculmorins were
more frequently found than CUL in Italian malting barley
samples156 and groundnuts from Cameroon.157 Besides factors
such as region, season, and sensitivity of the analytical method,
variations in the hydroxyculmorin proﬁle between individual
Fusarium species148 may account for these diﬀerences.
On the basis of available literature data, CUL seems to have
limited acute toxicity in mammals. However, due to its frequent
occurrence in cereal grains, the mode of action of this
mycotoxin should be elucidated and its potential interaction
with DON (and other trichothecenes) should be clariﬁed.
Butenolide. BUT (4-acetamido-4-hydroxy-2-butenoic acid
γ-lactone; Figure 7) is produced by several Fusarium species,

such as F. tricinctum, F. equiseti, F. graminearum, or
F. sporotrichioides. BUT was cytotoxic to diﬀerent established
mammalian cell lines126,158 and to primary cultures of neonatal
rat and murine cardiomyocytes.159,160 Minimal eﬀective
concentrations ranged from 1 to 25 μg/mL. In the case of
human chondrocytes, exposure to BUT concentrations ≥4 μg/
mL had a cytotoxic eﬀect, whereas lower concentrations (1−2
μg/mL) caused an increase in cell viability.161 ROS production
was shown to be a mechanism of BUT cytotoxicity in HepG2
cells.158 Furthermore, ROS production due to BUT exposure
caused lipid peroxidation159,161 and DNA damage160 in
mammalian cell cultures. BUT was shown to induce
mitochondrial dysfunction in rat cardiomyocytes, which may
trigger ROS production.162
The eﬀect of BUT exposure was investigated in rodents.
Intragastric administration of 10 or 20 mg BUT/kg BW to rats
for ≥7 weeks caused a loss of BW163 and injuries to the
myocardium159 and liver.163 Markers of oxidative stress and

oxidative damage were increased in myocardium, liver, and
serum.159,163 Exposure of mice to 0.5 mg BUT/mL drinking
water for 3 weeks (corresponds to consumption of ∼2.95 mg
BUT/day) had no eﬀect on BW.164 BUT showed adverse
eﬀects on the growth and development of rat embryos in vitro
when applied at a concentration of ≥1.25 mg/L, whereas no
such eﬀects were observed at 0.625 mg/L.165 In chicken
embryos, oxidative damage was detected in the myocardium,166
liver, and kidney167 after in ovo injection of a single dose of
BUT (10−100 μg). The toxicity of BUT was investigated in a
small number of cows. Application of ≥1.5 mg BUT to the skin
of heifers induced local inﬂammatory encrusted lesions.168
Intramuscular administration of 1.1 g BUT/day for 3 months,
oral administration of 4.5 g BUT/day for 2 months,168 or
intramuscular administration of 3.8 mg BUT/kg BW/day for 90
days169 caused weight loss and necrosis of the tail tip. Oral
administration of 39 and 68 mg/kg BW was lethal to steers
within 3 and 2 days, respectively.170 Oral administration of 31
mg/kg BW/day for 46 days caused weight loss and esophageal
and gastric ulcers.170 BUT was detected in 52% of samples (n =
83) of feed and feed raw materials with median and maximum
concentrations of 23 and 1490 μg/kg, respectively.60
In summary, the toxicity of BUT was shown in vivo and the
compound’s potential to trigger oxidative stress and oxidative
damage has been reported. However, very high doses of BUT
were administered in previous studies, and it is unclear whether
BUT levels that can be reached by dietary exposure would also
cause adverse eﬀects. Additional experiments with lower
concentrations of BUT should be conducted in vivo to study
the long-term toxicity. Also, more occurrence data are needed.
Sterigmatocystin. STE (Figure 8) is a toxic precursor of
aﬂatoxins and structurally closely related to AfB1. Its toxic eﬀect

is mediated by its furofuran ring structure, which forms DNA
adducts after metabolic activation to an epoxide.171 STE is
produced by several fungal species within the genera Aspergillus,
Bipolaris, Emericella, Chaetomium, Botryotrichum, and Humicola
and the species Penicillium luteum. The main producers are
Aspergillus fungi, such as Aspergillus ﬂavus, A. parasiticus,
A. nidulans, and especially A. versicolor.172 The International
Agency of Research in Cancer classiﬁed STE as a group 2B
carcinogen (possibly carcinogenic to humans).173,174
Several studies could demonstrate that STE exerts genotoxic
and cytotoxic eﬀects on diﬀerent cell lines, for example, on
immortalized ovarian hamster cells (CHO-K1 cells) for which
an IC50 value of 12.5 ± 2.0 μM was detected after 72 h of
incubation.175 The IC50 value of STE was determined to be 7.3
μM by measuring the protein synthesis in liver hepatocellular
(HepG2) cells.176 In addition, Gao and co-workers investigated
the STE-induced DNA damage in HepG2 cells. They showed a
signiﬁcant dose-dependent increase of DNA strand breaks and
an increased intracellular ROS level if cells were exposed to 3
and 6 μM STE.177 Also, in the human lung adenocarcinoma cell

Figure 7. Chemical structure of butenolide.

Figure 8. Chemical structure of sterigmatocystin.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7057

line A549 (IC50 = 3.7 μM)178 and in transformed rat ﬁbroblasts
(AWRF)179 the cytotoxic eﬀect of STE was conﬁrmed.
Furthermore, Sun and co-workers reported the induction of
apoptosis in human peripheral blood lymphocytes incubated
with 2 μg/mL STE.180 STE induced DNA damage in primary
cultured human esophageal epithelial cells and immortalized
human esophageal epithelial cells (Het-1A) and caused G1 and
G2 phase arrest, respectively.181
Only a few recently conducted in vivo studies are available in
livestock, ﬁsh, and other animals. Whereas the eﬀects of STE
are similar to those of AfB1, its acute toxicity is much lower.
The LD50 values in rat are 120−166 and 60−85 mg/kg BW for
oral and ip administration, respectively.182 In monkeys, the
LD50 upon ip administration was determined to be 32 mg/kg
BW.183 Studies in ruminants found a negative eﬀect of a STE-
contaminated diet in cattle,184 but not in sheep.185 Dietary
exposure of dairy cows to ≤12 mg STE/kg feed caused bloody
diarrhea, reduced milk production, and was lethal in some
cases.184 In one pig trial with STE (30 μg/kg feed), negative
eﬀects, such as decreased feed intake, incidental diarrhea, and
necrotic alterations of the liver tissue, were observed.186 An
LD50 of 5−7 μg was reported for 5-day-old chicken embryos,
whereas 10 μg killed 90−100% of the embryos.187 Newly
hatched male chicks suﬀered from liver cirrhosis and altered
biochemical serum parameters after 7 weeks of dietary STE
exposure.188 Also, Sreemannarayana and co-workers described
the hepatotoxic and nephrotoxic impact of STE in
poultry.189−192 In carp and catﬁsh, the mycotoxin caused a
decrease in body growth and crude protein content.193 In Nile
tilapia ﬁsh, STE showed toxic and clastogenic eﬀects indicated
by a signiﬁcantly decreased BW and an increased incidence of
micronucleated red blood cells as well as chromosomal
aberrations in the kidney.194 STE down-regulated TNF-α,
interleukin (IL) 6, and IL-12 mRNA expression in isolated
peripheral blood mononuclear cells and peritoneal macrophage
cells of BALB/c mice, as well as TNF-α and IL-6 protein levels
in serum, suggesting an immunosuppressive eﬀect.195 Already a
single sc dose of STE (5 μg/g of BW) induced tumors in lung
and liver in newborn mice. Males seemed to show an even
higher susceptibility to those eﬀects than females.196
Chromosome aberrations were found in rat bone marrow
cells, when a dose of 31.2 mg STE/kg BW was injected ip.197
Scarce data about the natural occurrence of STE was
available until 2015. An external scientiﬁc report, assigned by
EFSA, presents data on 1259 samples of cereal grains, cereal
products, beer, and nuts collected over a period of 14 months
and analyzed for STE by a validated LC-MS/MS method.198
STE was detected in 10% of all samples, and the highest
frequency of contamination was seen in rice (21%) and oat
grains (22%) intended for human consumption. In beer,
peanut, and hazelnut samples, no STE concentration was
found.198 A review from 2010 reported the occurrence of STE
in green coﬀee beans, spices, nuts, beer, and the outer layer of
hard cheese colonized by A. versicolor.199
The EFSA Panel on Contaminants in the Food Chain
(CONTAM) published a scientiﬁc opinion in 2013 about the
presence of STE in food and feed without characterizing its
possible risk due to a lack of occurrence data and the need for
more sensitive analytical methods.172 With new data available
now, the Joint FAO/WHO Expert Committee on Food
Additives (JECFA) is currently working on a safety assessment
of STE proposed by the Codex Committee on Contaminants in
Foods.200 Due to the high STE concentrations used in vitro, a

clear relevance of the observed results is arguable. Thus, follow-
up experiments with low-dose STE treatments should be
conducted in vivo to study the long-term toxicity.
Emodin. EMO, an orange-red crystalline compound, is a
1,3,8-trihydroxy-6-methylanthraquinone (Figure 9) and is

produced by many Aspergillus species, but has also been
found to be produced by the genera Penicillium and
Talaromyces. Furthermore, it occurs as an active ingredient in
many plants,201,202 mainly in Fabaceae (Cassia spp.),
Polygonaceae (Rheum, Rumex, and Polygonum spp.), and
Rhamnaceae (Rhamnus and Ventilago spp.).203 In plants, the
anthraquinones (also aloe-emodin, chrysophanol, physcion or
rhein) are present as aglycones or as glycosides, such as
emodin-8-glucoside.203
In traditional Chinese medicine, EMO, mainly extracted
from the rhizome of Rheum palmatum L., is claimed to be a
therapeutic alternative for the prophylaxis and treatment of
various diseases and physical complaints. Several studies could
demonstrate that EMO inhibits the growth and proliferation of
cancer cells in vivo and in vitro.204,205 EMO induced apoptosis
in the human lung squamous carcinoma cell line CH27206 and
in murine leukemia WEHI-3 cells.207 It might also act as a
JAK2/STAT3 inhibitor, and it down-regulates the expression of
anti-apoptotic Mcl-1, thereby inducing cell death in multiple
myeloma cell lines.208 Antiviral activities were shown in
diﬀerent studies. EMO reduced the entry and replication of
Coxsakievirus in HEp-2 cells209 and inhibited hepatitis B virus
and Herpes simplex virus type 1 in vitro.210,211 EMO exerted
antibacterial activity against Bacillus subtilis (minimal inhibitory
concentration (MIC) = 28.9 μM), Staphylococcus aureus (MIC
= 14.4 μM)212 and methicillin-resistant or sensitive S. aureus
strains (MIC = 64 μg/mL; 237 μM).213 EMO was suggested to
be purgative, as it was shown to induce contraction of rat
isolated ileum tissue by releasing endogenous acetylcholine.214
EMO was also shown to possess antidiabetic activity by
activating the PPARγ215 or the AMPK216 signaling pathway.
Furthermore, EMO is considered to have anti-osteopor-
otic217,218 and antiallergic eﬀects.219 A recently published
review describes several positive eﬀects in detail.220
In contrast to EMO’s manifold beneﬁcial eﬀects, its negative
eﬀects have not been thoroughly studied. In human
mononuclear cells, EMO (3−30 μM) caused an immunosup-
pressive reaction by decreasing IL-1 and IL-2 production and
IL-2 receptor expression.221 It induced apoptosis in human T
cells222 and a human proximal tubular cell line (HK-2) via the
mitochondrial pathway.223 Furthermore, it inhibited the
stimulation of human peripheral blood mononuclear cells and
decreased the plasma IL-2 level.224 Published studies about the
genotoxicity of EMO are controversial. Mueller and co-workers
showed a genotoxic eﬀect using the micronucleus test and
mutation assay in mouse lymphoma L5178Y cells.225,226 Morita
and co-workers observed an induction of 6-thioguanine-
resistant mutation in the mouse mammary carcinoma cell line

Figure 9. Chemical structure of emodin.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7058

FM3A by 1−10 μg/mL EMO.227 Other researchers, however,
could not demonstrate a mutagenic eﬀect of EMO.228,229
Furthermore, Westendorf and co-workers discovered only a
slight response in the V79-HGPRT mutagenicity assay.230
In vivo, therapeutic eﬀects and toxicity were investigated
mainly in mice and rats. Two studies with collagen-induced
arthritis mice showed that EMO exerted an anti-inﬂammatory
eﬀect by inhibition of the NF-κB-pathway231 and reduction of
TNF-α and IL-6 in plasma, PGE2 production, and COX-2
protein expression.232 Following successful in vitro experi-
ments, Liu and co-workers conﬁrmed the inhibition of the
Coxsakie virus also in mice.209 Another mouse trial showed the
positive eﬀect of EMO on diesel exhaust particle induced lung
inﬂammation.233 EMO also reduced aldose reductase in the
lens of mice, which might play an important role in the
development of diabetes cataract.234 Negative eﬀects of EMO
(0.25−2 μg/mL; 0.92−7.4 μM) were demonstrated for the
survival rates and hatching success of zebra ﬁsh embryos.235
Furthermore, EMO induced nephrotoxicity in Sprague−
Dawley rats.236
Long-term dietary exposure to EMO (≤2.5 g/kg) for a
period of 2 years showed no carcinogenic eﬀect in rats or
mice.237 Similarly, neither prenatal mortality nor morphologic
or genotoxic changes were observed in EMO-fed mice and rats,
and NOAELs of 2500 and 850 mg/kg were deﬁned for mice
and rats, respectively.238
EMO is probably the most ubiquitous anthraquinone found
worldwide, especially in higher plant families but also in fungi,
trees, shrubs, lianas, and herbs.203 However, EMO receives little
attention as a fungal metabolite, and occurrence data of fungal
EMO in food and feed are scarce. One study reports the
detection of EMO in 89% of the analyzed 83 feed samples, at
median and maximum concentrations of 9.8 and 1570 μg/kg.60
Because of the high NOAEL in rodents238 and its use in
alternative human medicine for decades, the risk to animal or
human health from EMO is very likely marginal.
Mycophenolic Acid. MPA ((4E)-6-(4-hydroxy-6-methoxy-
7-methyl-3-oxo-5-phthalanyl)-4-methyl-4-hexenoic acid; Figure
10) was isolated already 1893 by an Italian physician,

Bartolomeo Gosio, from Penicillium brevicompactum (reviewed
by Bentley239). Besides its production by this fungus, MPA can
also be produced by P. roqueforti and P. carneum or by
Byssochlamys niveaa known producer of patulin.4 MPA holds
the distinction of being the ﬁrst ever puriﬁed antibiotic,239 but
it is widely used nowadays both as an immunosuppressive drug
for prophylaxis and treatment of organ rejection in trans-
plantations and as an antirheumatic drug.240 The main mode of
action of MPA is the speciﬁc inhibition of the enzyme inosine
monophosphate dehydrogenase, which is highly expressed in
proliferating cells such as T- and B-lymphocytes. The inhibition
of lymphocyte proliferation leads to immunosuppression.241
Although MPA is sometimes referred to as a mycotoxin, its
acute toxicity in animals is low, with LD50 values of around 450
mg/kg BW in rats and around 1900 mg/kg BW in mice after

oral application.242 The same authors assessed subacute toxicity
in rats and detected no adverse eﬀects at 10 mg/kg BW. At
higher doses growth retardation and anemia were diagnosed in
rats. Dogs died after 12 days when given daily doses of 80 mg/
kg BW and at lower doses showed anorexia, diarrhea, and
enteritis.242 MPA was not mutagenic, did not produce
chromosome aberrations, and was nontoxic when given to
nonhuman primates in higher doses than required for human
therapy.243 Used as an antirheumatic drug in human medicine,
daily doses of 1−3 g were generally tolerated, with gastro-
intestinal intolerance being the major observed side eﬀect.243 In
the following years, a multitude of eﬀects on diﬀerent cell types,
e.g. lymphocytes, monocytes, neutrophils as well as on
dendritic, mesangial, mast, vascular smooth muscle and
endothelial cells were discovered.244 The authors concluded
in their literature review that while rheumatic patients still
largely beneﬁted from MPA treatment, it holds the risk of
immune suppression, and therapy has been associated with
several, sometimes life-threatening, infections by viruses.
MPA has been shown to occur in various food and feedstuﬀs
in relatively high concentrations compared to other fungal
metabolites. Dry-cured ham and liver pa
̂

té were shown to be
good substrates for P. brevicompactum, and after inoculation,
levels of 11−14 mg/kg of MPA were found on the food
surfaces.245 All investigated Roquefort cheese samples were
found positive for MPA with levels up to 1.2 mg/kg, whereas
German blue-white soft cheese contained even 11 mg/kg.246 In
lower concentrations, MPA was also found to occur in red
wine247 and ginger.248 In silage, MPA was found in 32% of 233
investigated sampled at levels up to 35 mg/kg249 or even up to
48 mg/kg.250
Given the low toxicity of MPA, the only concern is its role in
unwanted immunosuppression.251 The extensive use of MPA in
medicine has indicated that these concerns are minimal.239
Levels in food do not even come close to therapeutic doses in
humans, so it is safe to assume that MPA should also not be of
any dietary concern. Although in feed the concentrations can be
higher, it was veriﬁed that 300 mg of MPA per sheep and day
from contaminated silage does not show any immunodepres-
sive eﬀects.252 In conclusion, MPA should not be termed a
“mycotoxin” as it is neither toxic nor does it show any other
adverse eﬀects on humans or animals at the concentration
levels found in food and feed.
Alternariol and Alternariol Monomethyl Ether. AOH
and its monomethyl ether AME are dibenzopyrone derivatives
(Figure 11), produced by a large variety of Alternaria spp.,
which mainly grow on vegetables, fruits, and cereals.253 In vitro,
AOH and AME were shown to be cytotoxic to HeLa cells, but
less toxic to Bacillus mycoides.254 AOH was furthermore shown
to be cytotoxic to human colon carcinoma cell lines, and
cytotoxicity was shown to be mediated by the activation of the
mitochondrial pathway of apoptosis.255 In Caco-2 cells, AOH

Figure 10. Chemical structure of mycophenolic acid.

Figure 11. Chemical structure of alternariol (R = H) and alternariol
monomethyl ether (R = CH3).

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7059

induced toxicity via cell cycle disruption, induction of
apoptosis/necrosis, and changes in mitochondrial membrane
potential.256 AOH and AME induced DNA strand breaks and
chromosomal aberration in diﬀerent cell lines.253,257−259 The
DNA damaging eﬀect of these compounds was traced back to
their binding to the minor groove of DNA and their action as
topoisomerase poisons.258,260 Furthermore, in mammalian cell
lines, AOH induced oxidative stress that resulted in oxidative
DNA damage in some studies,261,262 but not in others.263,264 In
line with these observations of DNA-damaging eﬀects, AOH
was shown to be mutagenic in mammalian cell lines.265,266
Mutagenicity assays in bacteria, on the other hand, produced
contradictory results. Both compounds were suggested to be
strongly mutagenic according to the B. subtilis rec-assay267 and
the E. coli ND160 reverse mutation assay.268−270 In the Ames
Salmonella test, the compounds were not or only weakly
mutagenic to strains TA98 and TA100,271−273 and AOH was
mutagenic to strain TA102.274 Eﬀects of both AOH and AME
on reproductive organs and the immune system have been
suggested on the basis of in vitro data. An estrogenic potential
of AOH was detected in a human endometrial adenocarcinoma
cell line; however, this potential was 10,000-fold lower
compared to the endogenous hormone 17β-estradiol.257
Interestingly, AOH showed synergistic estrogenic eﬀects in
combination with ZEN or its phase I metabolite α-
zearalenol.275 AOH and AME speciﬁcally inhibited progester-
one secretion in cultured porcine granulosa cells, which
suggests that contaminated food may aﬀect the reproductive
performance of pigs and other mammalian species.276 AOH
induced autophagy and senescence in murine macrophages277
and modiﬁed the morphology and cytokine secretion of murine
and human macrophages.278
With regard to in vivo toxicity, no mortality or teratogenic
eﬀects of AOH or AME on chicken embryos were detected at
doses up to 1 mg per egg.279 Chicks exposed to dietary AME at
levels up to 100 mg/kg feed for 4 weeks showed no mortality
or signiﬁcant loss in performance.279 No evidence of toxicity
was observed in rats when AME and AOH were fed for 21 days
at levels up to 24 and 39 mg/kg feed, respectively.280
Furthermore, no toxic eﬀects were observed when 3.75 mg of
AME was given to rats by oral gavage daily for 30 days.281 In a
study performed by Schuchardt and co-workers, AOH was
shown to be nontoxic to mice after single or repeated oral
application of doses as high as 2000 mg/kg BW.282 An in vivo
oral toxicokinetic study using 200 or 1000 mg/kg AOH
revealed low systemic absorption and rapid metabolization, and
thus it was concluded that target organ toxicity would likely be
restricted to the gastrointestinal tract.282 Furthermore, neither
the micronucleus assay applied to bone marrow nor the Comet
assay applied to liver tissue indicated systemic genotoxicity.
Whether AOH exhibits local genotoxicity in the gastrointestinal
tract remains to be determined.
The metabolism of AOH and AME was investigated in
diﬀerent animal species and cell lines, and diﬀerent classes of
metabolites have been observed. AOH and AME were shown
to be hydroxylated at their aromatic carbon atoms by
cytochrome P450, leading to the formation of catechols,282−284
and they were shown to be conjugated with glucuronic acid or
sulfate at their phenolic hydroxyl groups by UDP-glucuronosyl-
transferases and sulfotransferases, respectively.285,286 AME was
furthermore shown to be demethylated to AOH in vitro.284
Two studies investigated the fetotoxicity of AOH and AME.
AME was maternally toxic and fetotoxic when administered ip

to Syrian golden hamsters at a single dose of 200 mg/kg BW,
whereas no such eﬀects were observed at lower doses.281 In
mice, sc administration of AOH (100 mg/kg BW) resulted in
an increased percentage of dead and resorbed fetuses/litter and
runts/litter, whereas lower doses of AOH or AME did not elicit
a fetotoxic eﬀect.254 Interestingly, a synergistic fetotoxic eﬀect
of AOH and AME was observed when both substances were
administered at a dose of 25 mg/kg. AOH and AME were
suggested as etiological agents of human esophageal
cancer,253,287 but a causal relationship has not been
demonstrated so far. Interestingly, precancerous changes were
detected in the esophageal mucosa of mice fed 50 or 100 mg
AME/kg BW/day for 10 months.288
EFSA concluded in 2011 that the estimated chronic dietary
exposure of humans to AOH and AME exceeds the threshold
of toxicological concern for potentially genotoxic substances,
indicating a need to create additional toxicity data.289 According
to a survey of published occurrence data, AOH was detected in
31% and AME was detected in 6% of samples (n ∼ 300) of
European feed and agricultural commodities at maximum
concentrations of 1840 and 184 μg/kg for AOH and AME,
respectively.289 Also, in tomato-based foodstuﬀs collected in
Belgium, AOH and AME exceeded the threshold of
toxicological concern assigned by EFSA for potentially
genotoxic substances.290
In conclusion, AOH and AME showed genotoxic eﬀects in
vitro. However, administration of high doses that couldon
the basis of currently available data on the occurrence in food
and feedhardly be achieved in vivo did not elicit toxic or
genotoxic eﬀects in rodents. Nevertheless, given the genotoxic
potential, additional toxicity data should be generated. Eﬀects
of AOH and AME on reproductive organs and the immune
system were suggested by in vitro data and should be
investigated in vivo.
Tenuazonic Acid. TeA is a tetramic acid derivative (Figure
12) produced by Alternaria species that grow on vegetables,

fruits, and cereals253 and by plant pathogenic fungi such as
Phoma sorghina and Magnaporthe oryzae.291,292 The toxic eﬀect
of TeA is presumably based on its interference with protein
biosynthesis.293 The compound was nonmutagenic in the Ames
Salmonella test with or without metabolic activation or
nitrosylation.272−274
TeA was toxic to mice and rats, and LD50 values were
determined.294,295 Observed symptoms included diarrhea,
muscle tremor, and convulsion. For mice, LD50 values upon
oral administration were 81 mg/kg BW for females and 186 or
225 mg/kg BW for males. Upon iv administration the LD50
value was 115 mg/kg BW for females. Upon ip, sc, or iv
administration, LD50 values were 125−162 mg/kg BW for
males. For rats, LD50 values were 168 mg/kg BW for females
and 180 mg/kg BW for males upon oral administration and 157
mg/kg BW for females and 146 mg/kg BW for males upon iv
administration. After dietary exposure of mice to 25 mg TeA/
kg BW/day for 10 months precancerous changes were detected

Figure 12. Chemical structure of tenuazonic acid.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7060

in the esophageal mucosa.288 TeA was furthermore toxic to
beagle dogs (4 oral doses of 2.5 mg/kg BW per day) and
monkeys (daily oral dose of 89.6 mg/kg BW).294 Symptoms
included diarrhea, vomiting, and hemorrhages and were lethal
in most cases. In the case of monkeys, no adverse eﬀects were
detected for daily doses up to 48.8 mg/kg BW. TeA was toxic
to chickens. The compound was toxic in the chicken embryo
assay with an LD50 of 548 μg/egg, but it showed no teratogenic
eﬀect.279 For day-old chicks, the LD50 upon oral administration
was 37.5 mg/kg BW.296 Dietary exposure of broilers to 10 mg/
kg feed for 3 weeks or daily oral administration of ≥1.25 mg/kg
BW to broilers and layers caused macro- and microscopic
lesions in diﬀerent organs and led to decreased weight gain and
feed eﬃciency.296 In the same study, no adverse eﬀects were
detected for doses of 0.63 mg/kg BW. The toxicokinetics of
TeA were investigated in pigs and broilers.297 Oral or iv
administration of single doses of 0.05 mg TeA/kg BW had no
adverse eﬀects. The orally administered doses were completely
absorbed in both species and rapidly eliminated in pigs but not
in broilers.
According to a survey of published occurrence data
conducted by EFSA, TeA was detected in 15% of samples (n
∼ 300) of European feed and agricultural commodities at
concentrations between 500 and 4310 μg/kg.289 In a survey of
feed and feed raw materials, TeA was detected in 65% of
samples (n = 83) at median and maximum concentrations of 68
and 1980 μg/kg, respectively.60 Interestingly, high levels of TeA
(up to 1200 μg/kg) were found in Sorghum-based infant
food.298
In summary, administration of TeA was toxic to rodents and
chickens. In their scientiﬁc opinion from 2011, EFSA
concluded, on the basis of exposure estimates and due to a
lack of data indicating genotoxicity, that TeA is unlikely to be a
human health concern.289 However, more recent occurrence
data suggested that health risks cannot be excluded for infants
consuming larger quantities of Sorghum-based food and that
more data on the subject should be generated. EFSA
furthermore concluded that on the basis of in vivo toxicity
data296 and estimated exposure, adverse health eﬀects cannot
fully be ruled out for chickens.289 The recent ﬁnding that TeA
shows a high bioavailability and slow elimination in chickens
reinforces the notion that the toxin may be a cause for concern
for this species and that more data on the subject should be
collected.
■ CONCLUSIONS
Fungal spoilage of agricultural goods is accompanied by the
formation of mycotoxins, which pose a signiﬁcant health risk to
humans and livestock. As climatic conditions favor the growth
of diﬀerent fungi (and hence the formation of diﬀerent toxins)
in diﬀerent parts of the world, some populations are more
aﬀected than others, but mycotoxins are a global challenge.
Without any doubt, aﬂatoxins, due to their high acute and
chronic toxicity, have the most severe impact on human and
animal life. In addition, several other mycotoxins including
trichothecenes, fumonisins, OTA, or ZEN are of high
toxicological signiﬁcance and have been extensively studied,
and regulations are in force in many countries with the aim of
ensuring tolerable levels in food and feed. Eﬃcient and reliable
analytical methods have been developed and are used to control
foodstuﬀs for their compliance with regulations. During the
past decade, advances inparticularly LC-MS basedmethods
used for food and feed analysis have been enormous with

regard to their detection limits, speed, and capability to
determine multiple compounds.299 Nowadays, certain modern
analytical methods are able to detect hundreds of diﬀerent
compounds, for instance, fungal metabolites,300 in a variety of
food and feed samples. Many of these metabolites are irrelevant
in terms of food and feed safety, but for the ﬁrst time in history
occurrence data can be collected on a large scale before the
toxicity of the detected fungal metabolites is (and can be)
assessed. Sticking to a basic principle in toxicology, sola dosis
facit venenum (the dose makes the poison), both toxicity and
occurrence data are indispensable for risk assessment. Instead
of prioritizing research on natural toxins based on the most
bioactive compounds, we might have reached a point where
assigning priorities based on actual occurrence data in
foodstuﬀs is at least conceivable.
Given the enormous number of bioactive fungal metabolites,
every attempt to provide exhaustive coverage of their potential
health impact is doomed to fail. As a limited number of
currently nonregulated compounds of fungal origin had to be
selected for this paper, it is plausible that equally or even more
important ones were missed. Still, our aim was to critically
scrutinize the impact of the emerging mycotoxins covered here
and to propose a ranking list for the research community to
prioritize research eﬀorts. The bottom of this list is clearly
reserved for EMO and MPA. Both compounds have already
been used in human medicine for decades with negligible side
eﬀects even at doses that cannot be reached due to ingestion of
contaminated food. CUL would probably rank above these
compounds on our imaginary list. Although levels of
occurrence can be high, the toxicity of this compound in
mammals seems to be marginal. Yet, the mode of action of this
substance has not been elucidated and, in addition, its frequent
co-occurrence with DON may be a cause for concern.
Therefore, it should be clariﬁed whether synergism with
trichothecenes, as hinted in a few studies, actually exists.
Arguably next on the list are ENNs and BEA. Whereas these
compounds are clearly toxic in vitro, most in vivo data indicate
limited toxicity. The EFSA risk assessment for these
compounds stated no immediate concern for human health,
but also highlighted the need for long-term studies to assess
potential chronic toxic eﬀects in vivo.59 Although BUT is toxic
in vivo and causes oxidative stress to cells, it is currently
unknown if the occurring levels are suﬃcient to cause adverse
eﬀects and toxicity studies with lower doses should be
conducted in animals. A similar picture can be drawn for
STE, as long-term in vivo studies at lower doses are currently
lacking that would be needed to assess chronic toxicity.
Whereas the Alternaria metabolites AOH and AME showed
genotoxic eﬀects in vitro, genotoxicity could so far not be
conﬁrmed in vivo, and further studies on this subject would be
warranted. Other eﬀects caused by these compounds, for
example, on the reproductive or immune system, should also be
clariﬁed using in vivo trials. In the case of FP and FA there is
clearly a need for more studies. Whereas FP levels in cereals can
be high and toxic eﬀects were seen on larvae and chicken
embryos, further animal studies are currently missing. Even less
is known about FA other than that it shows neurochemical
eﬀects and that it is possibly synergistic to co-occurring
mycotoxins such as DON or fumonisins. MON would probably
end up on the upper end of the ranking. Although its acute
toxicity and occurrence levels are unproblematic for humans,
the situation is diﬀerent for poultry. Furthermore, the
discrepancies between the outcomes of studies investigating

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7061

genotoxic eﬀects of MON should be clariﬁed as quickly as
possible. The situation is similar for TeA. Although toxic to
animals, EFSA concluded that this compound is likely of no
human health concern mainly due to its low average
occurrence.289 Considering recent studies, risk cannot be
excluded, though, both for chickens and for infants consuming
Sorghum-based food.
With regard to Alternaria metabolites, interestingly, AOH,
AME, and TeA made only a minor contribution to the
genotoxicity of certain A. alternata extracts, whereas a high
genotoxic activity was attributed to unidentiﬁed compounds.263
Also, altertoxins were shown to be mutagenic in vitro,273,301,302
and altertoxin I was detected in 42% of analyzed feed samples
(n = 83) in a recent survey.60 Altertoxins are just one example
that several fungal metabolites, not explicitly discussed in this
paper, show high toxic potential. Because of this, together with
the recently available information on frequent occurrence, more
data on the exposure of humans and animals as well as in vivo
studies clarifying possible toxic eﬀects are warranted.
■ ASSOCIATED CONTENT
*S Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acs.jafc.6b03413.
ChemDrawings (ZIP)

■ AUTHOR INFORMATION
Corresponding Author
*(F.B.) E-mail: franz.berthiller@boku.ac.at. Phone: +43 1
47654 97313.
Funding
We thank the Austrian Federal Ministry of Science, Research
and Economy, the Austrian National Foundation for Research,
Technology and Development and BIOMIN Holding GmbH
for funding the Christian Doppler Laboratory for Mycotoxin
Metabolism. The ﬁnancial support of the Austrian Research
Promotion Agency (FFG, Grant 84821) is greatly acknowl-
edged.
Notes
The authors declare no competing ﬁnancial interest.
■ ABBREVIATIONS USED
ABC ATP-binding cassette
AfB1 aﬂatoxin B1
AME alternariol monomethyl ether
AOH alternariol
BEA beauvericin
BUT butenolide
BW body weight
CC50 concentration resulting in 50% cell viability
CHEST chick embryotoxicity screening test
CONTAM Panel on Contaminants in the Food Chain
CUL culmorin
DON deoxynivalenol
EFSA European Food Safety Authority
EMO emodin
ENNs enniatins
FA fusaric acid
FB1 fumonisin B1
FP fusaproliferin
ip intraperitoneal
iv intravenous

IC50 half-maximal inhibitory concentration
IL interleukin
JECFA Joint FAO/WHO Expert Committee on Food
Additives
LC-MS liquid chromatography−mass spectrometry
LD50 dose that is lethal to 50% of test subjects
LOAEL lowest observed adverse eﬀect level
MIC minimal inhibitory concentration
MON moniliformin
MPA mycophenolic acid
NOAEL no observed adverse eﬀect level
OTA ochratoxin A
ROS reactive oxygen species
sc subcutaneous
STE sterigmatocystin
TeA tenuazonic acid
ZEN zearalenone

■ REFERENCES
(1) Dictionary of Natural Products v25.1; Taylor & Francis Group,
http://dnp.chemnetbase.com (accessed July 26, 2016).
(2) Keller, N. P.; Turner, G.; Bennett, J. W. Fungal secondary
metabolism − from biochemistry to genomics. Nat. Rev. Microbiol.
2005, 3, 937−947.
(3) Bennett, J. W. Mycotoxins, mycotoxicoses, mycotoxicology and
mycopathologia. Mycopathologia 1987, 100, 3−5.
(4) Frisvad, J. C.; Thrane, U.; Samson, R. A.; Pitt, J. I. Important
mycotoxins and the fungi which produce them. Adv. Exp. Med. Biol.
2006, 571, 3−31.
(5) Bennett, J. W.; Klich, M. Mycotoxins. Clin. Microbiol. Rev. 2003,
16, 497−516.
(6) Kensler, T. W.; Roebuck, B. D.; Wogan, G. N.; Groopman, J. D.
Aflatoxin: a 50-year odyssey of mechanistic and translational
toxicology. Toxicol. Sci. 2011, 120 (Suppl. 1), S28−S48.
(7) McCormick, S. P.; Stanley, A. M.; Stover, N. A.; Alexander, N. J.
Trichothecenes: from simple to complex mycotoxins. Toxins 2011, 3,
802−814.
(8) Stockmann-Juvala, H.; Savolainen, K. A review of the toxic effects
and mechanisms of action of fumonisin B1. Hum. Exp. Toxicol. 2008,
27, 799−809.
(9) Malir, F.; Ostry, V.; Pfohl-Leszkowicz, A.; Malir, J.; Toman, J.
Ochratoxin A, 50 years of research. Toxins 2016, 8, E191.
(10) Zinedine, A.; Soriano, J. M.; Molto,́ J. C.; Mañes, J. Review on
the toxicity, occurrence, metabolism, detoxification, regulations and
intake of zearalenone: an oestrogenic mycotoxin. Food Chem. Toxicol.
2007, 45, 1−18.
(11) van Egmond, H. P.; Schothorst, R. C.; Jonker, M. A. Regulations
relating to mycotoxins in food: perspectives in a global and European
context. Anal. Bioanal. Chem. 2007, 389, 147−157.
(12) Berthiller, F.; Brera, C.; Crews, C.; Iha, M. H.; Krska, R.;
Lattanzio, V. M. T.; MacDonald, S.; Malone, R. J.; Maragos, C.;
Solfrizzo, M.; Stroka, J.; Whitaker, T. B. Developments in mycotoxin
analysis: an update for 2013−2014. World Mycotoxin J. 2015, 8, 5−35.
(13) Berthiller, F.; Brera, C.; Crews, C.; Iha, M. H.; Krska, R.;
Lattanzio, V. M. T.; MacDonald, S.; Malone, R. J.; Maragos, C.;
Solfrizzo, M.; Stroka, J.; Whitaker, T. B. Developments in mycotoxin
analysis: an update for 2014−2015. World Mycotoxin J. 2016, 9, 5−30.
(14) Mogensen, J. M.; Frisvad, J. C.; Thrane, U.; Nielsen, K. F.
Production of fumonisin B2 and B4 by Aspergillus niger on grapes and
raisins. J. Agric. Food Chem. 2010, 58, 954−958.
(15) Battilani, P.; Toscano, P.; Van der Fels-Klerx, H. J.; Moretti, A.;
Camardo Leggieri, M.; Brera, C.; Rortais, A.; Goumperis, T.;
Robinson, T. Aflatoxin B1 contamination in maize in Europe increases
due to climate change. Sci. Rep. 2016, 6, 24328.
(16) Miraglia, M.; Marvin, H. J.; Kleter, G. A.; Battilani, P.; Brera, C.;
Coni, E.; Cubadda, F.; Croci, L.; De Santis, B.; Dekkers, S.; Filippi, L.;
Hutjes, R. W.; Noordam, M. Y.; Pisante, M.; Piva, G.; Prandini, A.;

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7062

Toti, L.; van den Born, G. J.; Vespermann, A. Climate change and food
safety: an emerging issue with special focus on Europe. Food Chem.
Toxicol. 2009, 47, 1009−1021.
(17) Varga, E.; Wiesenberger, G.; Hametner, C.; Ward, T. J.; Dong,
Y.; Schöfbeck, D.; McCormick, S.; Broz, K.; Stückler, R.;
Schuhmacher, R.; Krska, R.; Kistler, H. C.; Berthiller, F.; Adam, G.
New tricks of an old enemy: isolates of Fusarium graminearum produce
a type A trichothecene mycotoxin. Environ. Microbiol. 2015, 17, 2588−
2600.
(18) Berthiller, F.; Crews, C.; Dall’Asta, C.; De Saeger, S.; Haesaert,
G.; Karlovsky, P.; Oswald, I. P.; Seefelder, W.; Speijers, G.; Stroka, J.
Masked mycotoxins: a review. Mol. Nutr. Food Res. 2013, 57, 165−186.
(19) Rychlik, M.; Humpf, H. U.; Marko, D.; Da
̈

nicke, S.; Mally, A.;
Berthiller, F.; Klaffke, H.; Lorenz, N. Proposal of a comprehensive
definition of modified and other forms of mycotoxins including
“masked” mycotoxins. Mycotoxin Res. 2014, 30, 197−205.
(20) Jestoi, M. Emerging Fusarium mycotoxins fusaproliferin,
beauvericin, enniatins, and moniliformin − a review. Crit. Rev. Food
Sci. Nutr. 2008, 48, 21−49.
(21) Vaclavikova, M.; Malachova, A.; Veprikova, Z.; Dzuman, Z.;
Zachariasova, M.; Hajslova, J. ‘Emerging’ mycotoxins in cereals
processing chains: changes of enniatins during beer and bread making.
Food Chem. 2013, 136, 750−757.
(22) Sy-Cordero, A. A.; Pearce, C. J.; Oberlies, N. H. Revisiting the
enniatins: a review of their isolation, biosynthesis, structure
determination and biological activities. J. Antibiot. 2012, 65, 541−549.
(23) Kamyar, M.; Rawnduzi, P.; Studenik, C. R.; Kouri, K.;
Lemmens-Gruber, R. Investigation of the electrophysiological proper-
ties of enniatins. Arch. Biochem. Biophys. 2004, 429, 215−223.
(24) Dornetshuber, R.; Heffeter, P.; Kamyar, M. R.; Peterbauer, T.;
Berger, W.; Lemmens-Gruber, R. Enniatin exerts p53-dependent
cytostatic and p53-independent cytotoxic activities against human
cancer cells. Chem. Res. Toxicol. 2007, 20, 465−473.
(25) Fornelli, F.; Minervini, F.; Logrieco, A. Cytotoxicity of fungal
metabolites to lepidopteran (Spodoptera f rugiperda) cell line (SF-9). J.
Invertebr. Pathol. 2004, 85, 74−79.
(26) Ivanova, L.; Skjerve, E.; Eriksen, G. S.; Uhlig, S. Cytotoxicity of
enniatins A, A1, B, B1, B2 and B3 from Fusarium avenaceum. Toxicon
2006, 47, 868−876.
(27) Meca, G.; Font, G.; Ruiz, M. J. Comparative cytotoxicity study
of enniatins A, A(1), A(2), B, B(1), B(4) and J(3) on Caco-2 cells,
Hep-G(2) and HT-29. Food Chem. Toxicol. 2011, 49, 2464−2469.
(28) Wätjen, W.; Debbab, A.; Hohlfeld, A.; Chovolou, Y.;
Kampkotter, A.; Edrada, R. A.; Ebel, R.; Hakiki, A.; Mosaddak, M.;
Totzke, F.; Kubbutat, M. H.; Proksch, P. Enniatins A1, B and B1 from
an endophytic strain of Fusarium tricinctum induce apoptotic cell death
in H4IIE hepatoma cells accompanied by inhibition of ERK
phosphorylation. Mol. Nutr. Food Res. 2009, 53, 431−440.
(29) Hoornstra, D.; Andersson, M. A.; Mikkola, R.; Salkinoja-
Salonen, M. S. A new method for in vitro detection of microbially
produced mitochondrial toxins. Toxicol. In Vitro 2003, 17, 745−751.
(30) Behm, C.; Degen, G. H.; Follmann, W. The Fusarium toxin
enniatin B exerts no genotoxic activity, but pronounced cytotoxicity in
vitro. Mol. Nutr. Food Res. 2009, 53, 423−430.
(31) Dornetshuber, R.; Heffeter, P.; Lemmens-Gruber, R.; Elbling,
L.; Marko, D.; Micksche, M.; Berger, W. Oxidative stress and DNA
interactions are not involved in enniatin- and beauvericin-mediated
apoptosis induction. Mol. Nutr. Food Res. 2009, 53, 1112−1122.
(32) Tonshin, A. A.; Teplova, V. V.; Andersson, M. A.; Salkinoja-
Salonen, M. S. The Fusarium mycotoxins enniatins and beauvericin
cause mitochondrial dysfunction by affecting the mitochondrial
volume regulation, oxidative phosphorylation and ion homeostasis.
Toxicology 2010, 276, 49−57.
(33) Ivanova, L.; Egge-Jacobsen, W. M.; Solhaug, A.; Thoen, E.;
Faeste, C. K. Lysosomes as a possible target of enniatin B-induced
toxicity in Caco-2 cells. Chem. Res. Toxicol. 2012, 25, 1662−1674.
(34) Gammelsrud, A.; Solhaug, A.; Dendele, B.; Sandberg, W. J.;
Ivanova, L.; Kocbach Bolling, A.; Lagadic-Gossmann, D.; Refsnes, M.;
Becher, R.; Eriksen, G.; Holme, J. A. Enniatin B-induced cell death and

inflammatory responses in RAW 267.4 murine macrophages. Toxicol.
Appl. Pharmacol. 2012, 261, 74−87.
(35) Prosperini, A.; Juan-Garcia, A.; Font, G.; Ruiz, M. J. Reactive
oxygen species involvement in apoptosis and mitochondrial damage in
Caco-2 cells induced by enniatins A, A(1), B and B(1). Toxicol. Lett.
2013, 222, 36−44.
(36) Kolf-Clauw, M.; Sassahara, M.; Lucioli, J.; Rubira-Gerez, J.;
Alassane-Kpembi, I.; Lyazhri, F.; Borin, C.; Oswald, I. P. The emerging
mycotoxin, enniatin B1, down-modulates the gastrointestinal toxicity
of T-2 toxin in vitro on intestinal epithelial cells and ex vivo on
intestinal explants. Arch. Toxicol. 2013, 87, 2233−2241.
(37) Tomoda, H.; Huang, X. H.; Cao, J.; Nishida, H.; Nagao, R.;
Okuda, S.; Tanaka, H.; Omura, S.; Arai, H.; Inoue, K. Inhibition of
acyl-CoA:cholesterol acyltransferase activity by cyclodepsipeptide
antibiotics. J. Antibiot. 1992, 45, 1626−1632.
(38) Mereish, K. A.; Solow, R.; Bunner, D. L.; Fajer, A. B. Interaction
of cyclic peptides and depsipeptides with calmodulin. Pept. Res. 1990,
3, 233−237.
(39) Dornetshuber, R.; Heffeter, P.; Sulyok, M.; Schumacher, R.;
Chiba, P.; Kopp, S.; Koellensperger, G.; Micksche, M.; Lemmens-
Gruber, R.; Berger, W. Interactions between ABC-transport proteins
and the secondary Fusarium metabolites enniatin and beauvericin. Mol.
Nutr. Food Res. 2009, 53, 904−920.
(40) Ivanova, L.; Uhlig, S.; Eriksen, G.; Johannessen, L. Enniatin B1
is a substrate of intestinal P-glycoprotein, multidrug resistance-
associated protein 2 and breast cancer resistance protein. World
Mycotoxin J. 2010, 3, 271−281.
(41) Hiraga, K.; Yamamoto, S.; Fukuda, H.; Hamanaka, N.; Oda, K.
Enniatin has a new function as an inhibitor of Pdr5p, one of the ABC
transporters in Saccharomyces cerevisiae. Biochem. Biophys. Res.
Commun. 2005, 328, 1119−1125.
(42) Niimi, K.; Harding, D. R.; Holmes, A. R.; Lamping, E.; Niimi,
M.; Tyndall, J. D.; Cannon, R. D.; Monk, B. C. Specific interactions
between the Candida albicans ABC transporter Cdr1p ectodomain and
a D-octapeptide derivative inhibitor. Mol. Microbiol. 2012, 85, 747−
767.
(43) Bosch, U.; Mirocha, C. J.; Abbas, H. K.; di Menna, M. Toxicity
and toxin production by Fusarium isolates from New Zealand.
Mycopathologia 1989, 108, 73−79.
(44) Ga
̈

umann, E.; Naef-Roth, S.; Ettlinger, L. Zur Gewinnung von
Enniatinen aus dem Myzel verschiedener Fusarien. Phytopath. Z. 1950,
16, 289−299.
(45) Rodriguez-Carrasco, Y.; Heilos, D.; Richter, L.; Sussmuth, R. D.;
Heffeter, P.; Sulyok, M.; Kenner, L.; Berger, W.; Dornetshuber-Fleiss,
R. Mouse tissue distribution and persistence of the food-borne
fusariotoxins enniatin B and beauvericin. Toxicol. Lett. 2016, 247, 35−
44.
(46) Manyes, L.; Escriva, L.; Serrano, A. B.; Rodriguez-Carrasco, Y.;
Tolosa, J.; Meca, G.; Font, G. A preliminary study in Wistar rats with
enniatin A contaminated feed. Toxicol. Mech. Methods 2014, 24, 179−
190.
(47) Juan, C.; Manyes, L.; Font, G.; Juan-Garcia, A. Evaluation of
immunologic effect of enniatin A and quantitative determination in
feces, urine and serum on treated Wistar rats. Toxicon 2014, 87, 45−
53.
(48) McKee, T. C.; Bokesch, H. R.; McCormick, J. L.; Rashid, M. A.;
Spielvogel, D.; Gustafson, K. R.; Alavanja, M. M.; Cardelline, J. H.,
2nd; Boyd, M. R. Isolation and characterization of new anti-HIV and
cytotoxic leads from plants, marine, and microbial organisms. J. Nat.
Prod. 1997, 60, 431−438.
(49) CODA-CERVA (Centrum voor Onderzoek in Diergeneeskunde
en AgrochemieCentre d‘E
́

tude et de Recherches Vé

té

rinaires et
Agrochimiques). Carry-over of mycotoxins to animal products: a case
study poultry. Scientiﬁc Report 2011/2012; 2011/2012, pp 141−144,
available at www.coda-cerva.be/images/pdf/CODA_CERVA_
Scientiﬁc_Report_2011_2012.pdf.
(50) Jestoi, M.; Rokka, M.; Ja
̈

rvenpaä,̈ E.; Peltonen, K. Determination
of Fusarium mycotoxins beauvericin and enniatins (A, A1, B, B1) in

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7063

eggs of laying hens using liquid chromatography−tandem mass
spectrometry (LC−MS/MS). Food Chem. 2009, 115, 1120−1127.
(51) Taevernier, L.; Bracke, N.; Veryser, L.; Wynendaele, E.; Gevaert,
B.; Peremans, K.; Spiegeleer, B. Blood-brain barrier transport kinetics
of the cyclic depsipeptide mycotoxins beauvericin and enniatins.
Toxicol. Lett. 2016, 258, 175−184.
(52) Devreese, M.; Broekaert, N.; De Mil, T.; Fraeyman, S.; De
Backer, P.; Croubels, S. Pilot toxicokinetic study and absolute oral
bioavailability of the Fusarium mycotoxin enniatin B1 in pigs. Food
Chem. Toxicol. 2014, 63, 161−165.
(53) Faeste, C. K.; Ivanova, L.; Uhlig, S. In vitro metabolism of the
mycotoxin enniatin B in different species and cytochrome p450
enzyme phenotyping by chemical inhibitors. Drug Metab. Dispos. 2011,
39, 1768−1776.
(54) Ivanova, L.; Faeste, C. K.; Delezie, E.; Van Pamel, E.; Daeseleire,
E.; Callebaut, A.; Uhlig, S. Presence of enniatin B and its hepatic
metabolites in plasma and liver samples from broilers and eggs from
laying hens. World Mycotoxin J. 2014, 7, 167−175.
(55) Ivanova, L.; Faeste, C. K.; Uhlig, S. In vitro phase I metabolism
of the depsipeptide enniatin B. Anal. Bioanal. Chem. 2011, 400, 2889−
2901.
(56) Levy, D.; Bluzat, A.; Seigneuret, M.; Rigaud, J. L. Alkali cation
transport through liposomes by the antimicrobial fusafungine and its
constitutive enniatins. Biochem. Pharmacol. 1995, 50, 2105−2107.
(57) Lund, V. J.; Grouin, J. M.; Eccles, R.; Bouter, C.; Chabolle, F.
Efficacy of fusafungine in acute rhinopharyngitis: a pooled analysis.
Rhinology 2004, 42, 207−212.
(58) European Medicines Agency. http://www.ema.europa.eu/ema/
index.jsp?curl=pages/medicines/human/referrals/Fusafungine_for_
oromucosal_and_nasal_use/human_referral_prac_000054.jsp (ac-
cessed July 26, 2016).
(59) EFSA Panel on Contaminants in the Food Chain (CONTAM).
Scientiﬁc Opinion on the risks to human and animal health related to
the presence of beauvericin and enniatins in food and feed. EFSA J.
2014, 12, 3802.10.2903/j.efsa.2014.3802
(60) Streit, E.; Schwab, C.; Sulyok, M.; Naehrer, K.; Krska, R.;
Schatzmayr, G. Multi-mycotoxin screening reveals the occurrence of
139 different secondary metabolites in feed and feed ingredients.
Toxins 2013, 5, 504−523.
(61) Hu, L.; Rychlik, M. Occurrence of enniatins and beauvericin in
60 Chinese medicinal herbs. Food Addit. Contam., Part A 2014, 31,
1240−1245.
(62) Kouri, K.; Lemmens, M.; Lemmens-Gruber, R. Beauvericin-
induced channels in ventricular myocytes and liposomes. Biochim.
Biophys. Acta, Biomembr. 2003, 1609, 203−210.
(63) Kouri, K.; Duchen, M. R.; Lemmens-Gruber, R. Effects of
beauvericin on the metabolic state and ionic homeostasis of ventricular
myocytes of the guinea pig. Chem. Res. Toxicol. 2005, 18, 1661−1668.
(64) Zhan, J.; Burns, A. M.; Liu, M. X.; Faeth, S. H.; Gunatilaka, A. A.
Search for cell motility and angiogenesis inhibitors with potential
anticancer activity: beauvericin and other constituents of two
endophytic strains of Fusarium oxysporum. J. Nat. Prod. 2007, 70,
227−232.
(65) Dombrink-Kurtzman, M. A. Fumonisin and beauvericin induce
apoptosis in turkey peripheral blood lymphocytes. Mycopathologia
2003, 156, 357−364.
(66) Jow, G. M.; Chou, C. J.; Chen, B. F.; Tsai, J. H. Beauvericin
induces cytotoxic effects in human acute lymphoblastic leukemia cells
through cytochrome c release, caspase 3 activation: the causative role
of calcium. Cancer Lett. 2004, 216, 165−173.
(67) Klaric, M. S.; Rumora, L.; Ljubanovic, D.; Pepeljnjak, S.
Cytotoxicity and apoptosis induced by fumonisin B(1), beauvericin
and ochratoxin A in porcine kidney PK15 cells: effects of individual
and combined treatment. Arch. Toxicol. 2008, 82, 247−255.
(68) Lin, H. I.; Lee, Y. J.; Chen, B. F.; Tsai, M. C.; Lu, J. L.; Chou, C.
J.; Jow, G. M. Involvement of Bcl-2 family, cytochrome c and caspase 3
in induction of apoptosis by beauvericin in human non-small cell lung
cancer cells. Cancer Lett. 2005, 230, 248−259.

(69) Wa
̈

tjen, W.; Debbab, A.; Hohlfeld, A.; Chovolou, Y.; Proksch, P.
The mycotoxin beauvericin induces apoptotic cell death in H4IIE
hepatoma cells accompanied by an inhibition of NF-κB-activity and
modulation of MAP-kinases. Toxicol. Lett. 2014, 231, 9−16.
(70) Mallebrera, B.; Font, G.; Ruiz, M. J. Disturbance of antioxidant
capacity produced by beauvericin in CHO-K1 cells. Toxicol. Lett. 2014,
226, 337−342.
(71) Fotso, J.; Smith, J. S. Evaluation of beauvericin toxicity with the
bacterial bioluminescence assay and the Ames mutagenicity bioassay. J.
Food Sci. 2003, 68, 1938−1941.
(72) Klaric, M. S.; Darabos, D.; Rozgaj, R.; Kasuba, V.; Pepeljnjak, S.
Beauvericin and ochratoxin A genotoxicity evaluated using the alkaline
comet assay: single and combined genotoxic action. Arch. Toxicol.
2010, 84, 641−650.
(73) Ficheux, A. S.; Sibiril, Y.; Parent-Massin, D. Effects of
beauvericin, enniatin b and moniliformin on human dendritic cells
and macrophages: an in vitro study. Toxicon 2013, 71, 1−10.
(74) Mei, L.; Zhang, L.; Dai, R. An inhibition study of beauvericin on
human and rat cytochrome P450 enzymes and its pharmacokinetics in
rats. J. Enzyme Inhib. Med. Chem. 2009, 24, 753−762.
(75) Omura, S.; Koda, H.; Nishida, H. Hypolipemics containing
beauvericin as acylcoenzyme A cholesterol acyltransferase inhibitor.
Patent JP 89-16115019890623, 1991.
(76) Leitgeb, R.; Lew, H.; Wetscherek, W.; Böhm, J.; Quinz, A.
Influence of fusariotoxins on growing and slaughtering performance of
broilers. Die Bodenkultur 1999, 50, 57−66.
(77) Leitgeb, R.; Lew, H.; Khidr, R.; Böhm, J.; Zollitsch, W.; Wagner,
E. Influence of Furarium toxins on growth and carcass characteristics
of turkeys. Die Bodenkultur 2000, 51, 171−178.
(78) Leitgeb, R.; Raffaseder, C.; Ruckenbauer, P.; Lemmens, M.;
Bohm, J.; Wagner, E.; Krska, R.; Parich, A. Impact of Fusarium toxins
on growth and slaughter performance of broilers and turkeys.
Mycotoxin Res. 2003, 19, 180−184.
(79) Zollitsch, W.; Raffaseder, C.; Böhm, J.; Wagner, W.; Leitgeb, R.
Impact of the mycotoxins moniliformin and beauvericin on growth
and carcass traits of broilers. Wien. Tieraerztl. Mschr. 2003, 90, 238−
243.
(80) Jestoi, M.; Rokka, M.; Peltonen, K. An integrated sample
preparation to determine coccidiostats and emerging Fusarium-
mycotoxins in various poultry tissues with LC-MS/MS. Mol. Nutr.
Food Res. 2007, 51, 625−637.
(81) Cole, R. J.; Kirksey, J. W.; Cutler, H. G.; Doupnik, B. L.;
Peckham, J. C. Toxin from Fusarium moniliforme. Effects on plants and
animals. Science 1973, 4080, 1324−1326.
(82) Logrieco, A.; Mule, G.; Moretti, A.; Bottalico, A. Toxigenic
Fusarium species and mycotoxins associated with maize ear rot in
Europe. Eur. J. Plant Pathol. 2002, 108, 597−609.
(83) Desjardins, A. E.; Maragos, C. M.; Proctor, R. H. Maize ear rot
and moniliformin contamination by cryptic species of Fusarium
subglutinans. J. Agric. Food Chem. 2006, 54, 7383−7390.
(84) Kokkonen, M.; Ojala, L.; Parikka, P.; Jestoi, M. Mycotoxin
production of selected Fusarium species at different culture conditions.
Int. J. Food Microbiol. 2010, 143, 17−25.
(85) Ledoux, D. R.; Bermudez, A. J.; Rottinghaus, G. E.; Broomhead,
J.; Bennett, G. A. Effects of feeding Fusarium f ujikuroi culture material,
containing known levels of moniliformin, in young broiler chicks.
Poult. Sci. 1995, 74, 297−305.
(86) Marasas, W. F.; Thiel, P. G.; Sydenham, E. W.; Rabie, C. J.;
Lubben, A.; Nelson, P. E. Toxicity and moniliformin production by
four recently described species of Fusarium and two uncertain taxa.
Mycopathologia 1991, 113, 191−197.
(87) Leslie, J. F.; Zeller, K. A.; Lamprecht, S. C.; Rheeder, J. P.;
Marasas, W. F. O. Toxicity, pathogenicity, and genetic differentiation
of five species of Fusarium from sorghum and millet. Phytopathology
2005, 95, 275−283.
(88) Scauflaire, J.; Gourgue, M.; Callebaut, A.; Munaut, F. Fusarium
temperatum, a mycotoxin-producing pathogen of maize. Eur. J. Plant
Pathol. 2012, 133, 911−922.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7064

(89) Hallas-Moeller, M.; Nielsen, K. F.; Frisvad, J. C. Production of
the Fusarium mycotoxin moniliformin by Penicillium melanoconidium. J.
Agric. Food Chem. 2016, 64, 4505−4510.
(90) Thiel, P. G. A molecular mechanism for the toxic action of
moniliformin, a mycotoxin produced by Fusarium moniliforme.
Biochem. Pharmacol. 1978, 27, 483−486.
(91) Burka, L. T.; Doran, J.; Wilson, B. J. Enzyme inhibition and the
toxic action of moniliformin and other vinylogous α-ketoacids.
Biochem. Pharmacol. 1982, 31, 79−84.
(92) Gathercole, P. S.; Thiel, P. G.; Hofmeyr, J. H. S. Inhibition of
pyruvate dehydrogenase complex by moniliformin. Biochem. J. 1986,
233, 719−723.
(93) Pirrung, M. C.; Nauhaus, S. K.; Singh, B. Cofactor-directed,
time-dependent inhibition of thiamine enzymes by the fungal toxin
moniliformin. J. Org. Chem. 1996, 61, 2592−2593.
(94) Ficheux, A. S.; Sibiril, Y.; Le Garrec, R.; Parent-Massin, D. In
vitro myelotoxicity assessment of the emerging mycotoxins beauver-
icin, enniatin B and moniliformin on human hematopoietic
progenitors. Toxicon 2012, 59, 182−191.
(95) Cetin, Y.; Bullerman, L. B. Cytotoxicity of Fusarium mycotoxins
to mammalian cell cultures as determined by the MTT bioassay. Food
Chem. Toxicol. 2005, 43, 755−764.
(96) Wu, W.; Liu, T.; Vesonder, R. F. Comparative cytotoxicity of
fumonisin B1 and moniliformin in chicken primary cell cultures.
Mycopathologia 1995, 132, 111−116.
(97) Knasmueller, S.; Bresgen, N.; Kassie, F.; Mersch-Sundermann,
V.; Gelderblom, W.; Zoehrer, E.; Eckl, P. M. Genotoxic effects of three
Fusarium mycotoxins, fumonisin B1, moniliformin and vomitoxin in
bacteria and in primary cultures of rat hepatocytes. Mutat. Res., Genet.
Toxicol. Environ. Mutagen. 1997, 391, 39−48.
(98) Norred, W. P.; Plattner, R. D.; Vesonder, R. F.; Bacon, C. W.;
Voss, K. A. Effects of selected secondary metabolites of Fusarium
moniliforme on unscheduled synthesis of DNA by rat primary
hepatocytes. Food Chem. Toxicol. 1992, 30, 233−237.
(99) Celik, M.; Yilmaz, S.; Aksoy, H.; Unal, F.; Yuzbasioglu, D.;
Donbak, L. Evaluation of the genotoxicity of Fusarium mycotoxin
moniliformin in human peripheral blood lymphocytes. Environ. Mol.
Mutagen. 2009, 50, 431−434.
(100) Morgan, M. K.; Fitzgerald, S. D.; Rottinghaus, G. E.; Bursian,
S. J.; Aulerich, R. J. Toxic effects to mink of moniliformin extracted
from Fusarium f ujikuroi culture material. Vet. Hum. Toxicol. 1999, 41,
1−5.
(101) Allen, N. K.; Burmeister, H. R.; Weaver, G. A.; Mirocha, C. J.
Toxicity of dietary and intravenously administered moniliformin to
broiler chickens. Poult. Sci. 1981, 60, 1415−1417.
(102) Burmeister, H. R.; Ciegler, A.; Vesonder, R. F. Moniliformin, a
metabolite of Fusarium moniliforme NRRL 6322: purification and
toxicity. Appl. Environ. Microbiol. 1979, 37, 11−13.
(103) Kriek, N. P. J.; Marasas, W. F. O.; Steyn, P. S.; Van Rensburg,
S. J.; Steyn, M. Toxicity of a moniliformin-producing strain of
Fusarium moniliforme var. subglutinans isolated from maize. Food
Cosmet. Toxicol. 1977, 15, 579−587.
(104) Jonsson, M.; Jestoi, M.; Nathanail, A. V.; Kokkonen, U.-M.;
Anttila, M.; Koivisto, P.; Karhunen, P.; Peltonen, K. Application of
OECD Guideline 423 in assessing the acute oral toxicity of
moniliformin. Food Chem. Toxicol. 2013, 53, 27−32.
(105) Bermudez, A. J.; Ledoux, D. R.; Rottinghaus, G. E.; Bennett, G.
A. The individual and combined effects of the Fusarium mycotoxins
moniliformin and fumonisin B1 in turkeys. Avian Dis. 1997, 41, 304−
311.
(106) Li, Y. C.; Ledoux, D. R.; Bermudez, A. J.; Fritsche, K. L.;
Rottinghaus, G. E. The individual and combined effects of fumonisin
B1 and moniliformin on performance and selected immune
parameters in turkey poults. Poult. Sci. 2000, 79, 871−878.
(107) Kamyar, M. R.; Kouri, K.; Rawnduzi, P.; Studenik, C.;
Lemmens-Gruber, R. Effects of moniliformin in presence of
cyclohexadepsipeptides on isolated mammalian tissue and cells.
Toxicol. In Vitro 2006, 20, 1284−1291.

(108) Hietaniemi, V.; Ramo, S.; Yli-Mattila, T.; Jestoi, M.; Peltonen,
S.; Kartio, M.; Sievilainen, E.; Koivisto, T.; Parikka, P. Updated survey
of Fusarium species and toxins in Finnish cereal grains. Food Addit.
Contam., Part A 2016, 33, 831−848.
(109) Mohale, S.; Medina, A.; Rodriguez, A.; Sulyok, M.; Magan, N.
Mycotoxigenic fungi and mycotoxins associated with stored maize
from different regions of Lesotho. Mycotoxin Res. 2013, 29, 209−219.
(110) Scarpino, V.; Blandino, M.; Negre, M.; Reyneri, A.; Vanara, F.
Moniliformin analysis in maize samples from North-West Italy using
multifunctional clean-up columns and the LC-MS/MS detection
method. Food Addit. Contam., Part A 2013, 30, 876−884.
(111) Labuda, R.; Parich, A.; Vekiru, E.; Tancinova, D. Incidence of
fumonisins, moniliformin and Fusarium species in poultry feed
mixtures from Slovakia. Ann. Agric. Environ. Med. 2005, 12, 81−86.
(112) Battilani, P.; Costa, L. G.; Dossena, A.; Gullino, M. L.;
Marchelli, R.; Galaverna, G.; Pietri, A.; Dall’Asta, C.; Giorni, P.;
Spadaro, D.; Gualla, A. Chapter 4 − Moniliformin in food and feed.
Scientiﬁc Information on Mycotoxins and Natural Plant Toxicants.
EFSA Supporting Publ. 2009, EFSA-Q-2008-04995, 193−249.
(113) Peltonen, K.; Jestoi, M.; Eriksen, G. S. Health effects of
moniliformin: a poorly understood Fusarium mycotoxin. World
Mycotoxin J. 2010, 3, 403−414.
(114) Jonsson, M.; Atosuo, J.; Jestoi, M.; Nathanail, A. V.; Kokkonen,
U.-M.; Anttila, M.; Koivisto, P.; Lilius, E.-M.; Peltonen, K. Repeated
dose 28-day oral toxicity study of moniliformin in rats. Toxicol. Lett.
2015, 233, 38−44.
(115) European Food Safety Authority, http://registerofquestions.
efsa.europa.eu/raw-war/mandateLoader?mandate=M-2010-0312 (ac-
cessed July 21, 2016).
(116) Logrieco, A.; Moretti, A.; Fornelli, F.; Fogliano, V.; Ritieni, A.;
Caiaffa, M. F.; Randazzo, G.; Bottalico, A.; Macchia, L. Fusaproliferin
production by Fusarium subglutinans and its toxicity to Artemia salina,
SF-9 insect cells, and IARC/LCL 171 human B lymphocytes. Appl.
Environ. Microbiol. 1996, 62, 3378−3384.
(117) Prosperini, A.; Meca, G.; Font, G.; Ruiz, M. J. Study of the
cytotoxic activity of beauvericin and fusaproliferin and bioavailability in
vitro on Caco-2 cells. Food Chem. Toxicol. 2012, 50, 2356−2361.
(118) Ritieni, A.; Monti, S. M.; Randazzo, G.; Logrieco, A.; Moretti,
A.; Peluso, G.; Ferracane, R.; Fogliano, V. Teratogenic effects of
fusaproliferin on chicken. J. Agric. Food Chem. 1997, 45, 3039−3043.
(119) Pocsfalvi, G.; Ritieni, A.; Randazzo, G.; Dobo, A.; Malorni, A.
Interaction of fusarium mycotoxins, fusaproliferin and fumonisin B1,
with DNA studied by electrospray ionization mass spectrometry. J.
Agric. Food Chem. 2000, 48, 5795−5801.
(120) Santini, A.; Meca, G.; Uhlig, S.; Ritieni, A. Fusaproliferin,
beauvericin and enniatins: occurrence in food − a review. World
Mycotoxin J. 2012, 5, 71−81.
(121) Ritieni, A.; Moretti, A.; Logrieco, A.; Bottalico, A.; Randazzo,
G.; Monti, S. M.; Ferracane, R.; Fogliano, V. Occurrence of
fusaproliferin, fumonisin B1, and beauvericin in maize from Italy. J.
Agric. Food Chem. 1997, 45, 4011−4016.
(122) Bacon, C. W.; Porter, J. K.; Norred, W. P.; Leslie, J. F.
Production of fusaric acid by Fusarium species. Appl. Environ. Mircobiol
1996, 62, 4039−4043.
(123) Desjardins, A. E. Fusarium Mycotoxins − Chemistry, Genetics
and Biology; APS Press, 2006; 260 pp.
(124) Velasco, M.; Gilbert, C. A.; Rutledge, C. O.; McNay, J. L.
Antihypertensive effect of a dopamine beta hydroxylase inhibitor,
bupicomide: a comparison with hydralazine. Clin. Pharmacol. Ther.
1975, 18, 145−153.
(125) Fernandez-Pol, J. A.; Klos, D. J.; Hamilton, P. D. Cytotoxic
activity of fusaric acid on human adenocarcinoma cells in tissue
culture. Anticancer Res. 1993, 13, 57−64.
(126) Vesonder, R. F.; Gasdorf, H.; Peterson, R. E. Comparison of
the cytotoxicities of Fusarium metabolites and Alternaria metabolite
AAL-toxin to cultured mammalian cell lines. Arch. Environ. Contam.
Toxicol. 1993, 24, 473−477.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7065

(127) Abdul, N. S.; Nagiah, S.; Chuturgoon, A. A. Fusaric acid
induces mitochondrial stress in human hepatocellular carcinoma
(HepG2) cells. Toxicon 2016, 119, 336−344.
(128) May, H. D.; Wu, Q.; Blake, C. K. Effects of the Fusarium spp.
mycotoxins fusaric acid and deoxynivalenol on the growth of
Ruminococcus albus and Methanobrevibacter ruminantium. Can. J.
Microbiol. 2000, 46, 692−699.
(129) Boonman, N.; Prachya, S.; Boonmee, A.; Kittakoop, P.;
Wiyakrutta, S.; Sriubolmas, N.; Warit, S.; Dharmkrong-At
Chusattayanond, A. In vitro acanthamoebicidal activity of fusaric
acid and dehydrofusaric acid from an endophytic fungus Fusarium sp.
Tlau3. Planta Med. 2012, 78, 1562−1567.
(130) Dong, X.; Ling, N.; Wang, M.; Shen, Q.; Guo, S. Fusaric acid is
a crucial factor in the disturbance of leaf water imbalance in Fusarium-
infected banana plants. Plant Physiol. Biochem. 2012, 60, 171−179.
(131) Selim, M. E.; El-Gammal, N. A. Role of fusaric acid mycotoxin
in pathogensis process of tomato wilt disease caused by Fusarium
oxysporum. J. Bioprocess. Biotech. 2015, 5, 10.
(132) Singh, V. K.; Upadhyay, R. S. Fusaric acid induced cell death
and changes in oxidative metabolism of Solanum lycopersicum L. Bot.
Stud. 2014, 55, 1−11.
(133) Hidaka, H.; Nagatsu, T.; Takeya, K. Fusaric acid, a hypotensive
agent produced by fungi. J. Antibiot. 1969, 22, 228−230.
(134) Stack, B. C., Jr.; Ye, J.; Willis, R.; Hubbard, M.; Hendrickson,
H. P. Determination of oral bioavailability of fusaric acid in male
Sprague-Dawley rats. Drugs D 2014, 14, 139−145.
(135) Hidaka, H. Fusaric (5-butylpicolinic) acid, an inhibitor of
dopamine beta-hydroxylase, affects serotonin and noradrenalin. Nature
1971, 231, 54−55.
(136) Terasawa, F.; Ying, L. H.; Kameyama, M. The hypotensive
effect of fusaric acid: the results of long-term administration of fusaric
acid in elderly hypertensive patients. Jpn. Circ. J. 1976, 40, 1017−1023.
(137) Porter, J. K.; Bacon, C. W.; Wray, E. M.; Hagler, W. M., Jr.
Fusaric acid in Fusarium moniliforme cultures, corn, and feeds toxic to
livestock and the neurochemical effects in the brain and pineal gland of
rats. Nat. Toxins 1995, 3, 91−100.
(138) Smith, T. K.; MacDonald, E. J. Effect of fusaric acid on brain
regional neurochemistry and vomiting behaviour in swine. J. Anim. Sci.
1991, 69, 2044−2049.
(139) Smith, T. K.; McMillan, E. G.; Castillo, J. B. Effect of feeding
blends of Fusarium mycotoxin-contaminated grains containing
deoxynivalenol and fusaric acid on growth and feed consumption of
immature swine. J. Anim. Sci. 1997, 75, 2184−2191.
(140) Chu, Q.; Wu, W.; Smalley, E. B. Decreased cell-mediated
immunity and lack of skeletal problems in broiler chickens consuming
diets amended with fusaric acid. Avian Dis. 1993, 37, 863−867.
(141) Yin, E. S.; Rakhmankulova, M.; Kucera, K.; de Sena Filho, J. G.;
Portero, C. E.; Narva
́

ez-Trujillo, A.; Holley, S. A.; Strobel, S. A. Fusaric
acid induces a notochord malformation in zebrafish via copper
chelation. BioMetals 2015, 28, 783−789.
(142) Bacon, C. W.; Porter, J. K.; Norred, W. P. Toxic interaction of
fumonisins B1 and fusaric acid measured by injection into fertile
chicken egg. Mycopathologia 1995, 129, 29−35.
(143) Ruda, J. M.; Beus, K. S.; Hollenbeak, C. S.; Wilson, R. P.; Stack,
B. C. The effect of single agent oral fusaric acid (FA) on the growth of
subcutaneously xenografted SCC-1 cells in a nude mouse model.
Invest. New Drugs 2006, 24, 377−381.
(144) Smith, T. K.; Sousadias, M. G. Fusaric acid content of swine
feedstuffs. J. Agric. Food Chem. 1993, 41, 2296−2298.
(145) Pedersen, P. B.; Miller, J. D. The fungal metabolite culmorin
and related compounds. Nat. Toxins 1999, 7, 305−309.
(146) Strongman, D.; Miller, J.; Calhoun, L.; Findlay, J.; Whitney, N.
The biochemical basis for interference competition among some
lignicolous marine fungi. Bot. Mar. 1987, 30, 21−26.
(147) Miller, J. D.; MacKenzie, S. Secondary metabolites of Fusarium
venenatum strains with deletions in the Tri5 gene encoding trichodiene
synthetase. Mycologia 2000, 92, 764−771.
(148) Langseth, W.; Ghebremeskel, M.; Kosiak, B.; Kolsaker, P.;
Miller, D. Production of culmorin compounds and other secondary

metabolites by Fusarium culmorum and F. graminearum strains isolated
from Norwegian cereals. Mycopathologia 2001, 152, 23−34.
(149) Wang, Y.; Miller, J. Effects of Fusarium graminearum
metabolites on wheat tissue in relation to Fusarium head blight
resistance. J. Phytopathol. 1988, 122, 118−125.
(150) McCormick, S. Genetic control of Fusarium mycotoxins to
enhance food safety. Research project 421045, 2014 annual report;
http://www.ars.usda.gov/research/projects/projects.htm?ACCN_
NO=421045&fy=2014 (accessed July 19, 2016).
(151) Prelusky, D.; Hamilton, R.; Trenholm, H. Application of the
chick embryotoxicity bioassay for the evaluation of mycotoxin toxicity.
Microbiol., Aliments, Nutr. 1989, 7, 57−65.
(152) Dowd, P. F.; Miller, J. D.; Greenhalgh, R. Toxicity and
interactions of some Fusarium graminearum metabolites to caterpillars.
Mycologia 1989, 81, 646−650.
(153) Rotter, R.; Trenholm, H.; Prelusky, D.; Hartin, K.; Thompson,
B.; Miller, J. A preliminary examination of potential interactions
between deoxynivalenol (DON) and other selected Fusarium
metabolites in growing pigs. Can. Vet. J. 1992, 72, 107−116.
(154) Ghebremeskel, M.; Langseth, W. The occurrence of culmorin
and hydroxy-culmorins in cereals. Mycopathologia 2001, 152, 103−
108.
(155) Uhlig, S.; Eriksen, G. S.; Hofgaard, I. S.; Krska, R.; Beltra
́

n, E.;
Sulyok, M. Faces of a changing climate: semi-quantitative multi-
mycotoxin analysis of grain grown in exceptional climatic conditions in
Norway. Toxins 2013, 5, 1682−1697.
(156) Beccari, G.; Caproni, L.; Tini, F.; Uhlig, S.; Covarelli, L.
Presence of Fusarium species and other toxigenic fungi in malting
barley and multi-mycotoxin analysis by liquid chromatography-high
resolution mass spectrometry. J. Agric. Food Chem. 2016, 64, 4390−
4399.
(157) Abia, W. A.; Warth, B.; Sulyok, M.; Krska, R.; Tchana, A. N.;
Njobeh, P. B.; Dutton, M. F.; Moundipa, P. F. Determination of multi-
mycotoxin occurrence in cereals, nuts and their products in Cameroon
by liquid chromatography tandem mass spectrometry (LC-MS/MS).
Food Control 2013, 31, 438−453.
(158) Wang, Y. M.; Peng, S. Q.; Zhou, Q.; Wang, M. W.; Yan, C. H.;
Yang, H. Y.; Wang, G. Q. Depletion of intracellular glutathione
mediates butenolide-induced cytotoxicity in HepG2 cells. Toxicol. Lett.
2006, 164, 231−238.
(159) Liu, J. B.; Wang, Y. M.; Peng, S. Q.; Han, G.; Dong, Y. S.;
Yang, H. Y.; Yan, C. H.; Wang, G. Q. Toxic effects of Fusarium
mycotoxin butenolide on rat myocardium and primary culture of
cardiac myocytes. Toxicon 2007, 50, 357−364.
(160) Yang, H. Y.; Wang, Y. M.; Peng, S. Q. Metallothionein-I/II null
cardiomyocytes are sensitive to Fusarium mycotoxin butenolide-
induced cytotoxicity and oxidative DNA damage. Toxicon 2010, 55,
1291−1296.
(161) Shi, Z.; Cao, J.; Chen, J.; Li, S.; Zhang, Z.; Yang, B.; Peng, S.
Butenolide induced cytotoxicity by disturbing the prooxidant-
antioxidant balance, and antioxidants partly quench in human
chondrocytes. Toxicol. In Vitro 2009, 23, 99−104.
(162) Wang, Y. M.; Liu, J. B.; Peng, S. Q. Effects of Fusarium
mycotoxin butenolide on myocardial mitochondria in vitro. Toxicol.
Mech. Methods 2009, 19, 79−85.
(163) Wang, H. J.; Wang, Y. M.; Peng, S. Q. Repeated administration
of a Fusarium mycotoxin butenolide to rats induces hepatic lipid
peroxidation and antioxidant defense impairment. Food Chem. Toxicol.
2009, 47, 633−637.
(164) Burmeister, H. R.; Grove, M. D.; Kwolek, W. F. Moniliformin
and butenolide: effect on mice of high-level, long-term oral intake.
Appl. Environ. Microbiol. 1980, 40, 1142−1144.
(165) Guo, J.; Zhang, L. S.; Wang, Y. M.; Yan, C. H.; Huang, W. P.;
Wu, J.; Yuan, H. T.; Lin, B. W.; Shen, J. L.; Peng, S. Q. Study of
embryotoxicity of Fusarium mycotoxin butenolide using a whole rat
embryo culture model. Toxicol. In Vitro 2011, 25, 1727−1732.
(166) Wang, Y. M.; Wang, H. J.; Peng, S. Q. Lipid peroxidation and
antioxidant defense impairment in the hearts of chick embryos induced

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7066

by in ovo exposure to Fusarium mycotoxin butenolide. Toxicon 2008,
52, 781−786.
(167) Wang, Y. M.; Wang, H. J.; Peng, S. Q. In ovo exposure of a
Fusarium mycotoxin butenolide induces hepatic and renal oxidative
damage in chick embryos, and antioxidants provide protections.
Toxicol. In Vitro 2009, 23, 1354−1359.
(168) Kosuri, N. R.; Grove, M. D.; Yates, S. G.; Tallent, W. H.; Ellis,
J. J.; Wolff, I. A.; Nichols, R. E. Response of cattle to mycotoxins of
Fusarium tricinctum isolated from corn and fescue. J. Am. Vet. Med.
Assoc. 1970, 157, 938−940.
(169) Grove, M. D.; Yates, S. G.; Tallent, W. H.; Ellis, J. J.; Wolff, I.
A.; Kosuri, N. R.; Nichols, R. E. Mycotoxins produced by Fusarium
tricinctum as possible causes of cattle disease. J. Agric. Food Chem.
1970, 18, 734−736.
(170) Tookey, H. L.; Yates, S. G.; Ellis, J. J.; Grove, M. D.; Nichols,
R. E. Toxic effects of a butenolide mycotoxin and of Fusarium
tricinctum cultures in cattle. J. Am. Vet. Med. Assoc. 1972, 160, 1522−
1526.
(171) Wang, J.-S.; Groopman, J. D. DNA damage by mycotoxins.
Mutat. Res., Fundam. Mol. Mech. Mutagen. 1999, 424, 167−181.
(172) EFSA Panel on Contaminants in the Food Chain
(CONTAM). Scientiﬁc opinion on the risk for public and animal
health related to the presence of sterigmatocystin in food and feed.
EFSA J. 2013, 11, 3254.10.2903/j.efsa.2013.3254
(173) IARC. International Agency for Research on Cancer. Some
Naturally Occurring Substances; Monographs, International Agency on
for Research Cancer: Lyon, France, 1976; Vol. 10, pp 245−251.
(174) IARC. International Agency for Research on Cancer. Some
Naturally Occurring Substances; Monographs, International Agency on
for Research Cancer: Lyon, France, 1987; Vol. 10, p 72.
(175) Zouaoui, N.; Mallebrera, B.; Berrada, H.; Abid-Essefi, S.;
Bacha, H.; Ruiz, M. J. Cytotoxic effects induced by patulin,
sterigmatocystin and beauvericin on CHO-K1 cells. Food Chem.
Toxicol. 2016, 89, 92−103.
(176) Liu, Y.; Du, M.; Zhang, G. Proapoptotic activity of aflatoxin B1
and sterigmatocystin in HepG2 cells. Toxicol. Rep. 2014, 1, 1076−
1086.
(177) Gao, W.; Jiang, L.; Ge, L.; Chen, M.; Geng, C.; Yang, G.; Li,
Q.; Ji, F.; Yan, Q.; Zou, Y.; Zhong, L.; Liu, X. Sterigmatocystin-induced
oxidative DNA damage in human liver-derived cell line through
lysosomal damage. Toxicol. In Vitro 2015, 29, 1−7.
(178) Bünger, J.; Westphal, G.; Mönnich, A.; Hinnendahl, B.; Hallier,
E.; Müller, M. Cytotoxicity of occupationally and environmentally
relevant mycotoxins. Toxicology 2004, 202, 199−211.
(179) Ste
̆

tina, R. Induction of DNA single-strand breaks and DNA
synthesis inhibition in CHO and AWRF cells after exposure to
sterigmatocystin and penicillic acid. Folia Biol. 1986, 32, 406−413.
(180) Sun, X. M.; Zhang, X. H.; Wang, H. Y.; Cao, W. J.; Yan, X.;
Zuo, L. F.; Wang, J. L.; Wang, F. Effects of sterigmatocystin,
deoxynivalenol and aflatoxin G1 on apoptosis of human peripheral
blood lymphocytes in vitro. Biomed. Environ. Sci. 2002, 15, 145−152.
(181) Wang, J.; Huang, S.; Xing, L.; Cui, J.; Tian, Z.; Shen, H.; Jiang,
X.; Yan, X.; Wang, J.; Zhang, X. Sterigmatocystin induces G1 arrest in
primary human esophageal epithelial cells but induces G2 arrest in
immortalized cells: key mechanistic differences in these two models.
Arch. Toxicol. 2015, 89, 2015−2025.
(182) Purchase, I. F.; van der Watt, J. J. Acute toxicity of
sterigmatocystin to rats. Food Cosmet. Toxicol. 1969, 7, 135−139.
(183) Purchase, I. F.; van der Watt, J. J. The acute and chronic
toxicity of sterigmatocystin. In Mycotoxins in Human Health:
Proceedings of a Symposium Held in Pretoria; Purchase, I. F. H., Ed.;
MacMillan: London, UK, 1970; pp 209−213.
(184) Vesonder, R. F.; Horn, B. W. Sterigmatocystin in dairy cattle
feed contaminated with Aspergillus versicolor. Appl. Environ. Microbiol.
1985, 49, 234−235.
(185) Böhm, J.; Sayed, A. Investigation on toxic effects of
sterigmatocystin in sheep. Wien. Tieraerztl. Mschr. 1994, 81, 60−64.

(186) Kovalenko, A. V.; Soldatenko, N. A.; Fetisov, L. N.; Strel
́

tsov,
N. V. More accurate determination of the minimum allowable level of
sterigmatocystin in piglet feed. Russ. Agric. Sci. 2011, 37, 504−507.
(187) Schroeder, H. W.; Kelton, W. H. Production of sterigmato-
cystin by some species of the genus Aspergillus and its toxicity to
chicken embryos. Appl. Microbiol. 1975, 30, 589−591.
(188) Sayed, A. Clinical, Haematological and Biochemical Studies on the
Eﬀect of Sterigmatocystin in Sheep. Doctoral thesis, University of
Veterinary Medicine, Vienna, Austria, 1993.
(189) Sreemannarayana, O.; Frohlich, A. A.; Marquardt, R. R. Serum
biochemical studies in chicks injected intraperitoneally with
sterigmatocystin. Poult. Adviser 1986, 19, 53−58.
(190) Sreemannarayana, O.; Marquardt, R. R.; Frohlich, A. A.; Juck,
F. A. Some acute biochemical and pathological-changes in chicks after
oral-administration of sterigmatocystin. Int. J. Toxicol. 1986, 5, 275−
287.
(191) Sreemannarayana, O.; Frohlich, A. A.; Marquardt, R. R. Acute
toxicity of sterigmatocystin to chicks. Mycopathologia 1987, 97, 51−59.
(192) Sreemannarayana, O.; Frohlich, A. A.; Marquardt, R. R. Effects
of repeated intra-abdominal injections of sterigmatocystin on relative
organ weights, concentration of serum and liver constituents, and
histopathology of certain organs of the chick. Poult. Sci. 1988, 67,
502−509.
(193) Abdelhamid, A. M. Effects of sterigmatocystin contaminated
diets on fish performance. Arch. Tierernaehr. 1988, 38, 833−846.
(194) Abdel-Wahhab, M. A.; Hasanb, A. M.; Alya, S. E.; Mahrousb,
K. F. Adsorption of sterigmatocystin by montmorillonite and
inhibition of its genotoxicity in the Nile tilapia fish (Oreachromis
nilaticus). Mutat. Res., Genet. Toxicol. Environ. Mutagen. 2005, 582, 20−
27.
(195) Zhang, Y.; Yao, Z. G.; Wand, J.; Xing, L. X.; Xia, Y.; Zhang, X.
H. Effects of sterigmatocystin on TNF-α, IL-6 and IL-12 expression in
murine peripheral blood mononuclear cells and peritoneal macro-
phages in vivo. Mol. Med. Rep. 2012, 5, 1318−1322.
(196) Fujii, K.; Kurata, H.; Odashima, S.; Hatsuda, Y. Tumor
induction by a single subcutaneous injection of sterigmatocystin in
newborn mice. Cancer Res. 1976, 36 (5), 1615−1618.
(197) Ueda, N.; Fujie, K.; Gotoh-Mimura, K.; Chattopadhyay, S. C.;
Sugiyama, T. Acute cytogenetic effect of sterigmatocystin on rat bone-
marrow cells in vivo. Mutat. Res. Lett. 1984, 139, 203−206.
(198) Mol, H. G. J.; Pietri, A.; MacDonald, S. J.; Anagnostopoulos,
C.; Spanjer, S. Survey on sterigmatocystin in food. EFSA Supporting
Publ. 2015, EFSA-Q-2013-00280, EN−774.
(199) Versilovskis, A.; De Saeger, S. Sterigmatocystin: occurrence in
foodstuffs and analytical methods −an overview. Mol. Nutr. Food Res.
2010, 54 (1), 136−147.
(200) Joint FAO/WHO Food Standards Programme Codex
Committee on Contaminants. Ninth Session, Delhi, India, March
16−20, 2015.
(201) Wells, J. M.; Cole, R. J.; Kirksey, J. W. Emodin, a toxic
metabolite of Aspergillus wentii isolated from weevil-damaged chest-
nuts. Appl. Microbiol. 1975, 30, 26−28.
(202) Frisvad, J. C.; Larsen, T. O. Chemodiversity in the genus
Aspergillus. Appl. Microbiol. Biotechnol. 2015, 99, 7859−7877.
(203) Izhaki, I. Emodin − a secondary metabolite with multiple
ecological functions in higher plants. New Phytol. 2002, 155, 205−217.
(204) Huang, P. H.; Huang, C. Y.; Chen, M. C.; Lee, Y. T.; Yue, C.
H.; Wang, H. Y.; Lin, H. Emodin and aloe-emodin suppress breast
cancer cell proliferation through ERα inhibition. Evidence-Based
Complement. Alternat. Med. 2013, 2013, 376123.
(205) Wang, Q. J.; Cai, X. B.; Liu, M. H.; Tan, X. J.; Jing, X. B.
Apoptosis induced by emodin is associated with alterations of
intracellular acidification and reactive oxygen species in EC-109 cells.
Biochem. Cell Biol. 2010, 88, 767−774.
(206) Lee, H. Z. Effects and mechanisms of emodin on cell death in
human lung squamous cell carcinoma. Br. J. Pharmacol. 2001, 134,
11−20.
(207) Chang, Y. C.; Lai, T. Y.; Yu, C. S.; Chen, H. Y.; Yang, J. S.;
Chueh, F. S.; Lu, C. C.; Chiang, J. H.; Huang, W. W.; Ma, C. Y.;

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7067

Chung, J. G. Emodin induces apoptotic death in murine
myelomonocytic leukemia WEHI-3 cells in vitro and enhances
phagocytosis in leukemia mice in vivo. Evidence-Based Complement.
Alternat. Med. 2011, 2011, 1−13.
(208) Muto, A.; Hori, M.; Sasaki, Y.; Saitoh, A.; Yasuda, I.; Maekawa,
T.; Uchida, T.; Asakura, K.; Nakazato, T.; Kaneda, T.; Kizaki, M.;
Ikeda, Y.; Yoshida, T. Emodin has a cytotoxic activity against human
multiple myeloma as a Janus-activated kinase 2 inhibitor. Mol. Cancer
Ther. 2007, 6, 987−994.
(209) Liu, Z.; Wei, F.; Chen, L. J.; Xiong, H. R.; Liu, Y. Y.; Luo, F.;
Hou, W.; Xiao, H.; Yang, Z. Q. In vitro and in vivo studies of the
inhibitory effects of emodin isolated from Polygonum cuspidatum on
Coxsakie virus. Molecules 2013, 18, 11842−11858.
(210) Shuangsuo, D.; Zhengguo, Z.; Yunru, C.; Xin, Z.; Baofeng, W.;
Lichao, Y.; Yan’an, C. Inhibition of the replication of hepatitis B virus
in vitro by emodin. Med. Sci. Monit. 2006, 12, 302−306.
(211) Hsiang, C. Y.; Ho, T. Y. Emodin is a novel alkaline nuclease
inhibitor that suppresses herpes simplex virus type 1 yields in cell
cultures. Br. J. Pharmacol. 2008, 155, 227−235.
(212) Chukwujekwu, J. C.; Coombes, P. H.; Mulholland, D. A.; van
Staden, J. Emodin, an antibacterial anthraquinone from the roots of
Cassia occidentalis. S. Afr. J. Bot. 2006, 72, 295−297.
(213) Hatano, T.; Uebayashi, H.; Ito, H.; Shiota, S.; Tsuchiya, T.;
Yoshida, T. Phenolic constituents of cassia seeds and antibacterial
effect of some naphthalenes and anthraquinones on methicillin-
resistant Staphylococcus aureus. Chem. Pharm. Bull. 1999, 47, 1121−
1127.
(214) Ali, S.; Watson, M. S.; Osborne, R. H. The stimulant cathartic,
emodin, contracts the rat isolated ileum by triggering release of
endogenous acetylcholine. Auton. Autacoid Pharmacol. 2004, 24, 103−
105.
(215) Liu, Y.; Jia, L.; Liu, Z. C.; Zhang, H.; Zhang, P. J.; Wan, Q.;
Wang, R. Emodin ameliorated high-glucose induced mesangial P38
over-activation and hypocontractility via activating PPARγ. Exp. Mol.
Med. 2009, 41, 648−655.
(216) Chen, Z.; Zhang, L.; Yi, J.; Yang, Z.; Zhang, Z.; Li, Z.
Promotion of adiponectin multimerization by emodin: a novel AMPK
activator with PPARγ-agonist activity. J. Cell. Biochem. 2012, 113,
3547−3558.
(217) Kang, D. M.; Yoon, K. H.; Kim, J. Y.; Oh, J. M.; Lee, M.; Jung,
S. T.; Juhng, S. K.; Lee, Y. H. CT imaging biomarker for evaluation of
emodin as a potential drug on LPS-mediated osteoporosis mice. Acad.
Radiol. 2014, 21, 457−462.
(218) Kim, J. Y.; Cheon, Y. H.; Kwak, S. C.; Baek, J. M.; Yoon, K. H.;
Lee, M. S.; Oh, J. Emodin regulates bone remodeling by inhibiting
osteoclastogenesis and stimulating osteoblast formation. J. Bone Miner.
Res. 2014, 29, 1541−1553.
(219) Wang, W.; Zhou, Q.; Liu, L.; Zou, K. Anti-allergic activity of
emodin on IgE-mediated activation in RBL-2H3 cells. Pharmacol. Rep.
2012, 64, 1216−1222.
(220) Dong, X.; Fu, J.; Yin, X.; Cao, S.; Li, X.; Lin, L.; Huyiligeqi; Ni,
J. Emodin: a review of its pharmacology, toxicity and pharmacoki-
netics. Phytother. Res. 2016, 30, 1207−1218.
(221) Huang, H. C.; Chang, J. H.; Tung, S. F.; Wu, R. T.; Foegh, M.
L.; Chu, S. H. Immunosuppressive effect of emodin, a free radical
generator. Eur. J. Pharmacol. 1992, 211, 359−364.
(222) Qu, K.; Shen, N. Y.; Xu, X. S.; Su, H. B.; Wei, J. C.; Tai, M. H.;
Meng, F. D.; Zhou, L.; Zhang, Y. L.; Liu, C. Emodin induces human T
cell apoptosis in vitro by ROS-mediated endoplasmic reticulum stress
and mitochondrial dysfunction. Acta Pharmacol. Sin. 2013, 34, 1217−
1228.
(223) Wang, C.; Jiang, Z.; Yao, J.; Wu, X.; Sun, L.; Liu, C.; Duan, W.;
Yan, M.; Sun, L.; Liu, J.; Zhang, L. Participation of cathepsin B in
emodin-induced apoptosis in HK-2 cells. Toxicol. Lett. 2008, 181,
196−204.
(224) Liu, Y. X.; Shen, N. Y.; Liu, C.; Lv, Y. Immunosuppressive
effects of emodin: an in vivo and in vitro study. Transplant. Proc. 2009,
41, 1837−1839.

(225) Mueller, S. O.; Eckert, I.; Lutz, W. K.; Stopper, H. Genotoxicity
of the laxative drug components emodin, aloe-emodin and danthron in
mammalian cells: topoisomerase II mediated? Mutat. Res., Genet.
Toxicol. Test. 1996, 371, 165−173.
(226) Mueller, S. O.; Schmitt, M.; Dekant, W.; Stopper, H.; Schlatter,
J.; Schreier, P.; Lutz, W. K. Occurrence of emodin, chrysophanol and
physcion in vegetables, herbs and liquors. Genotoxicity and anti-
genotoxicity of the anthraquinones and of the whole plants. Food
Chem. Toxicol. 1999, 37, 481−491.
(227) Morita, H.; Umeda, M.; Masuda, T.; Ueno, Y. Cytotoxic and
mutagenic effects of emodin on cultured mouse carcinoma FM3A
cells. Mutat. Res., Genet. Toxicol. Test. 1988, 204, 329−332.
(228) Mengs, U.; Krumbiegel, G.; Völkner, W. Lack of emodin
genotoxicity in the mouse micronucleus assay. Mutat. Res., Genet.
Toxicol. Environ. Mutagen. 1997, 393, 289−293.
(229) Bruggemann, I. M.; van der Hoeven, J. C. M. Lack of activity of
the bacterial mutagen emodin in HGPRT and SCE assay with V79
Chinese hamster cells. Mutat. Res., Genet. Toxicol. Test. 1984, 138,
219−224.
(230) Westendorf, J.; Marquardt, H.; Poginsky, B.; Dominiak, M.;
Schmidt, J.; Marquardt, H. Genotoxicity of naturally occurring
hydroxyanthraquinones. Mutat. Res., Genet. Toxicol. Test. 1990, 240,
1−12.
(231) Hwang, J. K.; Noh, E. M.; Moon, S. J.; Kim, J. M.; Kwon, K. B.;
Park, B. H.; You, Y. O.; Hwang, B. M.; Kim, H. J.; Kim, B. S.; Lee, S. J.;
Kim, J. S.; Lee, Y. R. Emodin suppresses inflammatory responses and
joint destruction in collagen-induced arthritic mice. Rheumatology
2013, 52, 1583−1591.
(232) Zhu, X.; Zeng, K.; Qiu, Y.; Yan, F.; Lin, C. Therapeutic effect
of emodin on collagen-induced arthritis in mice. Inflammation 2013,
36, 1253−1259.
(233) Nemmar, A.; Salam, S.; Yuvaraju, P.; Beegam, S.; Alis, B. H.
Emodin mitigates diesel exhaust particles-induced increase in airway
resistance, inflammation and oxidative stress in mice. Respir. Physiol.
Neurobiol. 2015, 215, 51−57.
(234) Chang, K. C.; Li, L.; Sanborn, T. M.; Shieh, B.; Lenhart, P.;
Ammar, D.; LaBarbera, D. V.; Petrash, J. M. Characterization of
emodin as a therapeutic agent for diabetic cataract. J. Nat. Prod. 2016,
79, 1439−1444.
(235) He, Q.; Liu, K.; Wang, S.; Hou, H.; Yuan, Y.; Wang, X.
Toxicity induced by emodin on zebrafish embryos. Drug Chem.
Toxicol. 2012, 36, 149−154.
(236) Wang, C.; Dai, X.; Liu, H.; Yi, H.; Zhou, D.; Liu, C.; Ma, M.;
Jiang, Z.; Zhang, L. Involvement of PPARγ in emodin-induced HK-2
cell apoptosis. Toxicol. In Vitro 2015, 29, 228−233.
(237) NTP Toxicology and carcinogenesis studies of emodin (CAS
No. 518-82-1) feed studies in F344/N rats and B6C3F1 mice. Natl.
Toxicol. Program Tech. Rep. Ser. 2001, 493, 1−278.
(238) Jahnke, G. D.; Price, C. J.; Marr, M. C.; Myers, C. B.; George, J.
D. Developmental toxicity evaluation of emodin in rats and mice. Birth
Defects Res., Part B 2004, 71, 89−101.
(239) Bentley, R. Mycophenolic acid: a one hundred year odyssey
from antibiotic to immunosuppressant. Chem. Rev. 2000, 100, 3801−
3826.
(240) Cholewinski, G.; Malachowska-Ugarte, M.; Dzierzbicka, K.
The chemistry of mycophenolic acid − synthesis and modifications
towards desired biological activity. Curr. Med. Chem. 2010, 17, 1926−
1941.
(241) Allison, A. C.; Kowalski, W. J.; Muller, C. D.; Eugui, E. M.
Mechanisms of action of mycophenolic acid. Ann. N. Y. Acad. Sci.
1993, 696, 63−87.
(242) Sweeney, M. J.; Gerzon, K.; Harris, P. N.; Holmes, R. E.;
Poore, G. A.; Williams, R. H. Experimental antitumor activity and
preclinical toxicology of mycophenolic acid. Cancer Res. 1972, 32,
1795−1802.
(243) Allison, A. C.; Eugui, E. M. Immunosuppressive and other
effects of mycophenolic acid and an ester prodrug, mycophenolate
mofetil. Immunol. Rev. 1993, 136, 5−28.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7068

(244) Zizzo, G.; De Santis, M.; Ferraccioli, G. F. Mycophenolic acid
in rheumatology: mechanisms of action and severe adverse events.
Reumatismo 2010, 62, 91−100.
(245) Sørensen, L. M.; Nielsen, K. F.; Jacobsen, T.; Koch, A. G.;
Nielsen, P. V.; Frisvad, J. C. Determination of mycophenolic acid in
meat products using mixed mode reversed phase-anion exchange
clean-up and liquid chromatography-high-resolution mass spectrom-
etry. J. Chromatogr A 2008, 1205, 103−108.
(246) Usleber, E.; Dade, M.; Schneider, E.; Dietrich, R.; Bauer, J.;
Ma
̈

rtlbauer, E. Enzyme immunoassay for mycophenolic acid in milk
and cheese. J. Agric. Food Chem. 2008, 56, 6857−6862.
(247) Meyer, K.; Müller, T.; Dietrich, R.; Ma
̈

rtlbauer, E.; Bauer, J.
Nachweis und Vorkommen von Mykophenolsa
̈

ure und Citrinin in
Rotwein. Mycotoxin Res. 2001, 17 (Suppl. 2), 160−164.
(248) Overy, D. P.; Frisvad, J. C. Mycotoxin production and
postharvest storage rot of ginger (Zingiber of f icinale) by Penicillium
brevicompactum. J. Food Prot. 2005, 68, 607−609.
(249) Schneweis, I.; Meyer, K.; Hörmansdorfer, S.; Bauer, J.
Mycophenolic acid in silage. Appl. Environ. Microbiol. 2000, 66,
3639−3641.
(250) Gallo, A.; Bertuzzi, T.; Giuberti, G.; Moschini, M.; Bruschi, S.;
Cerioli, C.; Masoero, F. New assessment based on the use of principal
factor analysis to investigate corn silage quality from nutritional traits,
fermentation end products and mycotoxins. J. Sci. Food Agric. 2016, 96,
437−448.
(251) Kirby, B.; Yates, V. M. Mycophenolate mofetil for psoriasis. Br.
J. Dermatol. 1998, 139, 357.
(252) Dzidic, A.; Meyer, H. H. D.; Bauer, J.; Pfaffl, M. W. Long-term
effects of mycophenolic acid on the immunoglobulin and inflamma-
tory marker-gene expression in sheep white blood cells. Mycotoxin Res.
2010, 26, 235−240.
(253) Ostry, V. Alternaria mycotoxins: an overview of chemical
characterization, producers, toxicity, analysis and occurrence in
foodstuffs. World Mycotoxin J. 2008, 1, 175−188.
(254) Pero, R. W.; Posner, H.; Blois, M.; Harvan, D.; Spalding, J. W.
Toxicity of metabolites produced by the “Alternaria”. Environ. Health
Perspect. 1973, 4, 87−94.
(255) Bensassi, F.; Gallerne, C.; Sharaf El Dein, O.; Hajlaoui, M. R.;
Bacha, H.; Lemaire, C. Cell death induced by the Alternaria mycotoxin
alternariol. Toxicol. In Vitro 2012, 26, 915−923.
(256) Fernandez-Blanco, C.; Juan-Garcia, A.; Juan, C.; Font, G.; Ruiz,
M. J. Alternariol induce toxicity via cell death and mitochondrial
damage on Caco-2 cells. Food Chem. Toxicol. 2016, 88, 32−39.
(257) Lehmann, L.; Wagner, J.; Metzler, M. Estrogenic and
clastogenic potential of the mycotoxin alternariol in cultured
mammalian cells. Food Chem. Toxicol. 2006, 44, 398−408.
(258) Marko, D. Mechanisms of the genotoxic eﬀect of Alternaria
toxins. In 29th Mycotoxin Workshop; Gesellschaft für Mykotoxin-
Forschung: Stuttgart-Fellbach, Germany, 2007; p 48.
(259) Pfeiffer, E.; Eschbach, S.; Metzler, M. Alternaria toxins: DNA
strand-breaking activity in mammalian cells in vitro. Mycotoxin Res.
2007, 23, 152−157.
(260) Fehr, M.; Pahlke, G.; Fritz, J.; Christensen, M. O.; Boege, F.;
Altemoller, M.; Podlech, J.; Marko, D. Alternariol acts as a
topoisomerase poison, preferentially affecting the IIα isoform. Mol.
Nutr. Food Res. 2009, 53, 441−451.
(261) Fernandez-Blanco, C.; Font, G.; Ruiz, M. J. Oxidative DNA
damage and disturbance of antioxidant capacity by alternariol in Caco-
2 cells. Toxicol. Lett. 2015, 235, 61−66.
(262) Solhaug, A.; Vines, L. L.; Ivanova, L.; Spilsberg, B.; Holme, J.
A.; Pestka, J.; Collins, A.; Eriksen, G. S. Mechanisms involved in
alternariol-induced cell cycle arrest. Mutat. Res., Fundam. Mol. Mech.
Mutagen. 2012, 738−739, 1−11.
(263) Schwarz, C.; Kreutzer, M.; Marko, D. Minor contribution of
alternariol, alternariol monomethyl ether and tenuazonic acid to the
genotoxic properties of extracts from Alternaria alternata infested rice.
Toxicol. Lett. 2012, 214, 46−52.
(264) Tiessen, C.; Fehr, M.; Schwarz, C.; Baechler, S.; Domnanich,
K.; Bottler, U.; Pahlke, G.; Marko, D. Modulation of the cellular redox

status by the Alternaria toxins alternariol and alternariol monomethyl
ether. Toxicol. Lett. 2013, 216, 23−30.
(265) Brugger, E. M.; Wagner, J.; Schumacher, D. M.; Koch, K.;
Podlech, J.; Metzler, M.; Lehmann, L. Mutagenicity of the mycotoxin
alternariol in cultured mammalian cells. Toxicol. Lett. 2006, 164, 221−
230.
(266) Fleck, S. C.; Burkhardt, B.; Pfeiffer, E.; Metzler, M. Alternaria
toxins: altertoxin II is a much stronger mutagen and DNA strand
breaking mycotoxin than alternariol and its methyl ether in cultured
mammalian cells. Toxicol. Lett. 2012, 214, 27−32.
(267) Kada, T.; Sadaie, Y.; Sakamoto, Y. I. Bacillus subtilis repair test.
In Handbook of Mutagenicity Test Procedures, 2nd ed.; Kilbey, B. J.,
Legator, M., Nichols, W., Ramel, C., Eds.; Elsevier: Amsterdam, The
Netherlands, 1984; pp 13−31.
(268) An, Y.; Zhao, T.; Miao, J.; Liu, G.; Zheng, Y.; Xu, Y.; VanEtten,
R. L. Isolation, identification, and mutagenicity of alternariol
monomethyl ether. J. Agric. Food Chem. 1989, 37, 1341−1343.
(269) Yu-Hui, A.; Tian-Zheng, Z.; Jian, M.; Ying-Zhong, Z.; You-
Mei, X.; Gui-Ting, L.; VanEtten, R. The studies of isolation,
identification and mutagenicity of alternariol monomethyl ether.
Acta Acad. Med. Henan 1986, 21, 204−207.
(270) Zhen, Y. Z.; Xu, Y. M.; Liu, G. T.; Miao, J.; Xing, Y. D.; Zheng,
Q. L.; Ma, Y. F.; Su, T.; Wang, X. L.; Ruan, L. R. Mutagenicity of
Alternaria alternata and Penicillium cyclopium isolated from grains in an
area of high incidence of oesophageal cancer −Linxian, China. IARC
Sci. Publ. 1991, 253−257.
(271) Davis, V. M.; Stack, M. E. Evaluation of alternariol and
alternariol methyl ether for mutagenic activity in Salmonella
typhimurium. Appl. Environ. Microbiol. 1994, 60, 3901−3902.
(272) Schrader, T. J.; Cherry, W.; Soper, K.; Langlois, I.; Vijay, H. M.
Examination of Alternaria alternata mutagenicity and effects of
nitrosylation using the Ames Salmonella test. Teratog., Carcinog.,
Mutagen. 2001, 21, 261−274.
(273) Scott, P. M.; Stoltz, D. R. Mutagens produced by Alternaria
alternata. Mutat. Res., Genet. Toxicol. Test. 1980, 78, 33−40.
(274) Schrader, T. J.; Cherry, W.; Soper, K.; Langlois, I. Further
examination of the effects of nitrosylation on Alternaria alternata
mycotoxin mutagenicity in vitro. Mutat. Res., Genet. Toxicol. Environ.
Mutagen. 2006, 606, 61−71.
(275) Vejdovszky, K.; Hahn, K.; Braun, D.; Warth, B.; Marko, D.
Synergistic estrogenic effects of Fusarium and Alternaria mycotoxins in
vitro. Arch. Toxicol. 2016, DOI: 10.1007/s00204-016-1795-7.
(276) Tiemann, U.; Tomek, W.; Schneider, F.; Muller, M.; Pohland,
R.; Vanselow, J. The mycotoxins alternariol and alternariol methyl
ether negatively affect progesterone synthesis in porcine granulosa
cells in vitro. Toxicol. Lett. 2009, 186, 139−145.
(277) Solhaug, A.; Torgersen, M. L.; Holme, J. A.; Lagadic-
Gossmann, D.; Eriksen, G. S. Autophagy and senescence, stress
responses induced by the DNA-damaging mycotoxin alternariol.
Toxicology 2014, 326, 119−129.
(278) Solhaug, A.; Wisbech, C.; Christoffersen, T. E.; Hult, L. O.;
Lea, T.; Eriksen, G. S.; Holme, J. A. The mycotoxin alternariol induces
DNA damage and modify macrophage phenotype and inflammatory
responses. Toxicol. Lett. 2015, 239, 9−21.
(279) Griffin, G. F.; Chu, F. S. Toxicity of the Alternaria metabolites
alternariol, alternariol methyl ether, altenuene, and tenuazonic acid in
the chicken embryo assay. Appl. Environ. Microbiol. 1983, 46, 1420−
1422.
(280) Sauer, D. B.; Seitz, L. M.; Burroughs, R.; Mohr, H. E.; West, J.
L.; Milleret, R. J.; Anthony, H. D. Toxicity of Alternaria metabolites
found in weathered sorghum grain at harvest. J. Agric. Food Chem.
1978, 26, 1380−1393.
(281) Pollock, G. A.; DiSabatino, C. E.; Heimsch, R. C.; Hilbelink, D.
R. The subchronic toxicity and teratogenicity of alternariol
monomethyl ether produced by Alternaria solani. Food Chem. Toxicol.
1982, 20, 899−902.
(282) Schuchardt, S.; Ziemann, C.; Hansen, T. Combined
toxicokinetic and in vivo genotoxicity study on Alternaria toxins.
EFSA Supporting Publ. 2014, EFSA-Q-2012-00433, EN−679.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 7052−7070

7069

(283) Burkhardt, B.; Wittenauer, J.; Pfeiffer, E.; Schauer, U. M.;
Metzler, M. Oxidative metabolism of the mycotoxins alternariol and
alternariol-9-methyl ether in precision-cut rat liver slices in vitro. Mol.
Nutr. Food Res. 2011, 55, 1079−1086.
(284) Pfeiffer, E.; Schebb, N. H.; Podlech, J.; Metzler, M. Novel
oxidative in vitro metabolites of the mycotoxins alternariol and
alternariol methyl ether. Mol. Nutr. Food Res. 2007, 51, 307−316.
(285) Burkhardt, B.; Pfeiffer, E.; Metzler, M. Absorption and
metabolism of the mycotoxins alternariol and alternariol-9-methyl
ether in Caco-2 cells in vitro. Mycotoxin Res. 2009, 25, 149−157.
(286) Pfeiffer, E.; Schmit, C.; Burkhardt, B.; Altemöller, M.; Podlech,
J.; Metzler, M. Glucuronidation of the mycotoxins alternariol and
alternariol-9-methyl ether in vitro: chemical structures of glucuronides
and activities of human UDP-glucuronosyltransferase isoforms.
Mycotoxin Res. 2009, 25, 3−10.
(287) Liu, G. T.; Qian, Y. Z.; Zhang, P.; Dong, W. H.; Qi, Y. M.;
Guo, H. T. Etiological role of Alternaria alternata in human esophageal
cancer. Chin Med. J. (Engl.) 1992, 105, 394−400.
(288) Yekeler, H.; Bitmis, K.; Ozcelik, N.; Doymaz, M. Z.; Calta, M.
Analysis of toxic effects of Alternaria toxins on esophagus of mice by
light and electron microscopy. Toxicol. Pathol. 2001, 29, 492−497.
(289) EFSA Panel on Contaminants in the Food Chain
(CONTAM). Scientiﬁc Opinion on the risks for animal and public
health related to the presence of Alternaria toxins in feed and food.
EFSA J. 2011, 9, 2407.10.2903/j.efsa.2011.2407
(290) Walravens, J.; Mikula, H.; Rychlik, M.; Asam, S.; Devos, T.;
Njumbe Ediage, E.; Diana Di Mavungu, J.; Jacxsens, L.; Van
Landschoot, A.; Vanhaecke, L.; De Saeger, S. Validated UPLC-MS/
MS methods to quantitate free and conjugated Alternaria toxins in
commercially available tomato products and fruit and vegetable juices
in Belgium. J. Agric. Food Chem. 2016, 64, 5101−5109.
(291) Steyn, P. S.; Rabie, C. J. Characterization of magnesium and
calcium tenuazonate from Phoma sorghina. Phytochemistry 1976, 15,
1977−1979.
(292) Iwasaki, S.; Muro, H.; Nozoe, S.; Okuda, S.; Sato, Z. Isolation
of 3,4-dihydro-3,4,8-trihydroxy-2(2H)-naphthalenone and tenuazonic
acid from Pyricularia oryzae Cavara. Tetrahedron Lett. 1972, 13, 13−16.
(293) Shigeura, H. T.; Gordon, C. N. The biological activity of
tenuazonic acid. Biochemistry 1963, 2, 1132−1137.
(294) Smith, E. R.; Frederickson, T. N.; Hadidian, Z. Toxic effects of
the sodium and the N,N′-dibenzylethylenediamine salts of tenuazonic
acid (NSC-525816 and NSC-82260). Cancer Chemother. Rep. 1968,
52, 579−585.
(295) Woody, M. A.; Chu, F. S. Toxicology of Alternaria mycotoxins.
In Alternaria: Biology, Plant Diseases and Metabolites; Chelkowsky, J.,
Visconti, A., Eds.; Elsevier: Amsterdam, The Netherlands, 1992; pp
409−434.
(296) Giambrone, J. J.; Davis, N. D.; Diener, U. L. Effect of
tenuazonic acid on young chickens. Poult. Sci. 1978, 57, 1554−1558.
(297) Fraeyman, S.; Devreese, M.; Broekaert, N.; De Mil, T.;
Antonissen, G.; De Baere, S.; De Backer, P.; Rychlik, M.; Croubels, S.
Quantitative determination of tenuazonic acid in pig and broiler
chicken plasma by LC-MS/MS and its comparative toxicokinetics. J.
Agric. Food Chem. 2015, 63, 8560−8567.
(298) Asam, S.; Rychlik, M. Potential health hazards due to the
occurrence of the mycotoxin tenuazonic acid in infant food. Eur. Food
Res. Technol. 2013, 236, 491−497.
(299) Malik, A. K.; Blasco, C.; Pico,́ Y. Liquid chromatography-mass
spectrometry in food safety. J. Chromatogr A 2010, 1217, 4018−4040.
(300) Malachova,́ A.; Sulyok, M.; Beltrá

n, E.; Berthiller, F.; Krska, R.
Optimization and validation of a quantitative liquid chromatography-
tandem mass spectrometric method covering 295 bacterial and fungal
metabolites including all regulated mycotoxins in four model food
matrices. J. Chromatogr A 2014, 1362, 145−156.
(301) Davis, V. M.; Stack, M. E. Mutagenicity of stemphyltoxin III, a
metabolite of Alternaria alternata. Appl. Environ. Microbiol. 1991, 57,
180−182.

(302) Stack, M. E.; Prival, M. J. Mutagenicity of the Alternaria
metabolites altertoxins I, II, and III. Appl. Environ. Microbiol. 1986, 52,
718−722.

Journal of Agricultural and Food Chemistry Review

DOI: 10.1021/acs.jafc.6b03413
J. Agric. Food Chem. 2017, 65, 70

Author
Recent Posts

Dr. Raymond Oenbrink

Dr. Raymond Oenbrink DO is board certified by the American Osteopathic Board of Family Physicians (AOBFP) and the American Board of Addiction Medicine (ABAM). He specializes in complex and chronic illnesses such as Chronic Inflammatory Response Syndrome (CIRS).

Latest posts by Dr. Raymond Oenbrink (see all)

COVID UPDATE: What is the truth? - 2022-11-08
Pathologist Speaks Out About COVID Jab Effects - 2022-07-04
A Massive Spike in Disability is Most Likely Due to a Wave of Vaccine Injuries - 2022-06-30