Next Article in Journal
Introduction to the Toxins Special Issue on Plant Toxins
Next Article in Special Issue
Delay of the Onset of Puberty in Female Rats by Prepubertal Exposure to T-2 Toxin
Previous Article in Journal
Honeybee Venom Proteome Profile of Queens and Winter Bees as Determined by a Mass Spectrometric Approach
Previous Article in Special Issue
High Sensitivity of Aged Mice to Deoxynivalenol (Vomitoxin)-Induced Anorexia Corresponds to Elevated Proinflammatory Cytokine and Satiety Hormone Responses
Open AccessReview

Effects of Mycotoxins on Mucosal Microbial Infection and Related Pathogenesis

by Seong-Hwan Park 1,2,†, Dongwook Kim 3,†, Juil Kim 1,2 and Yuseok Moon 1,4,*
Laboratory of Mucosal Exposome and Biomodulation, Department of Biomedical Sciences, Pusan National University School of Medicine, Yangsan 50612, Korea
Research Institute for Basic Sciences and Medical Research Institute, Pusan National University, Busan 46241, Korea
National Institute of Animal Science, RDA, Wanju 55365, Korea
Immunoregulatory Therapeutics Group in Brain Busan 21 Project, Busan 46241, Korea
Author to whom correspondence should be addressed.
These authors contributed equally to this work.
Academic Editor: Jiujiang Yu
Toxins 2015, 7(11), 4484-4502;
Received: 4 September 2015 / Revised: 27 October 2015 / Accepted: 28 October 2015 / Published: 30 October 2015
(This article belongs to the Special Issue Mycotoxins and Human Diseases 2015)


Mycotoxins are fungal secondary metabolites detected in many agricultural commodities and water-damaged indoor environments. Susceptibility to mucosal infectious diseases is closely associated with immune dysfunction caused by mycotoxin exposure in humans and other animals. Many mycotoxins suppress immune function by decreasing the proliferation of activated lymphocytes, impairing phagocytic function of macrophages, and suppressing cytokine production, but some induce hypersensitive responses in different dose regimes. The present review describes various mycotoxin responses to infectious pathogens that trigger mucosa-associated diseases in the gastrointestinal and respiratory tracts of humans and other animals. In particular, it focuses on the effects of mycotoxin exposure on invasion, pathogen clearance, the production of cytokines and immunoglobulins, and the prognostic implications of interactions between infectious pathogens and mycotoxin exposure.
Keywords: Mycotoxins; microbial infection; mucosal pathogenesis Mycotoxins; microbial infection; mucosal pathogenesis

1. Introduction

Mycotoxins are natural, low-molecular-weight secondary fungal metabolites that are detected in various agricultural commodities and humid indoor environments, such as water-damaged buildings [1,2,3,4]. Exposure to mycotoxins in human and animals affects the host immune responses to infectious agents. Various immune-related organs or tissues are impaired by mycotoxins, which alters the susceptibility to the pathogens. Moreover, mycotoxins themselves alter the virulence of the infectious pathogens, leading to changes in the toxicity and invasiveness of the microbes in diverse organs or immune cells [5,6]. The most well-known crosstalk between mycotoxicosis and infection is the aflatoxin B1 exposure with the chronic hepatitis B virus infection, which has synergistic effects on the risk of hepatocellular carcinoma [7,8]. The main sites of mycotoxin exposure are the mucosal epithelia in the gut and airways with the underlying mucosal lymphoid tissues. In particular, the present review examines the interaction between mycotoxins and mucosa-associated pathogenic bacteria or viruses; this interaction exerts detrimental effects on target organs or cells by altering physiological or immunological conditions. The purpose of this review is to provide insights into the crucial roles of mycotoxins (including deoxynivalenol, fumonisin, T-2 toxin, aflatoxin, and ochratoxin) in the pathogenesis of mucosa-associated pathogens and their related mucosal disorders.

2. Mycotoxin Exposure and Altered Host Immune Responses

Various mycotoxins affect immune-related organs and cells, and influence host defenses against infectious agents and related microbial toxins [9]. Many previous studies have shown that aflatoxins suppress immune functions, particularly cell-mediated immune responses [10,11]. For instance, high levels of aflatoxin B1 (AFB1)-albumin adducts change T-cell phenotypes and reduce the percentage of B cells in human immunodeficiency virus-positive individuals [12]. Moreover, exposure to ochratoxin A (OTA) results in reductions in the sizes of crucial immune organs in various species, including the thymus [13] and bursa of Fabricius with advanced atrophy of bursal follicles [14,15,16]. OTA induces necrosis of the germinal center of the spleen and lymph nodes in rats and dogs. Mice exposed to OTA have a decreased number of hematopoietic stem cells [13] and total white blood cells [17]. OTA-exposed mice have fewer splenic T lymphocytes and mature CD4+ cells with more immature double-positive CD4+/CD8+ cells [18]. OTA exposure also decreases IL-2 production and IL-2 receptor expression in porcine and human T lymphocyte populations and subpopulations [19,20]. OTA and citrinin as the nephrotoxic mycotoxins are the etiological factor of the fatal human kidney disease and show synergistic lethal responses and suppression of lymphocyte proliferation in a synergistic way [21,22]. In addition to lymphocytes, embryonic exposure to AFB1 impairs the functions of phagocytes such as macrophages and neutrophils, via the depression of phagocytic potential, inhibition of antiviral activity, and reduction in chemotactic responses [23,24,25]. AFB1 also interferes with the innate immunity of macrophages by suppressing tumor necrosis factor-α (TNF- α), interleukin (IL)-1, and IL-6, resulting in the disruption of pulmonary and systemic host defenses [11,26]. Exposure to fumonisin B1 (FB1) also alters the morphology and functions of macrophages in chickens, leading to increased susceptibility to bacterial infections [27]. Additionally, natural killer cell-mediated cytolysis and resistance to infectious agents are also reduced by exposure to aflatoxins [28,29,30,31] and deoxynivalenol (DON) [32].
Humoral immune responses are also affected by mycotoxin exposure. FB1-producing Fusarium moniliforme can lead to deficiencies in antibody titers in the chicken immune system [33]. Moreover, the white blood cell population and antibody production are reduced in DON-exposed mice compared with unexposed mice [34,35]. In particular, IgM and delayed-type hypersensitivity responses to infectious bacteria are significantly suppressed. However, serum IgA levels increase after exposure to DON, leading to mesangial deposition of the IgA-immune complex, although serum IgM levels decrease [34]. Other animals, such as chicks and pigs, exhibit increased antibody responses after exposure to DON [36,37]. Depending on the dose regime, the immune response can be differentially regulated by mycotoxins. Low-dose exposure to DON or other type B trichothecene mycotoxins, including nivalenol, 15-acetyl DON, and 3-acetyl DON, induces chemokine production in human or mouse intestinal epithelial cells (IECs) [38,39], IL-2 production in human lymphocytes, and pro-inflammatory cytokine production, including IL-8, IL-6, and TNF-α in human macrophages [40]. Therefore, human and animal immune responses are altered under exposure to mycotoxins, which leads to impaired pathological damages to mycotoxin-exposed tissues or organs.

3. Interaction between Mycotoxins and Pathogenic Infections in the GI Tract

3.1. Relationship between Mycotoxin Exposure and Salmonella Infection

Salmonella is a genus of flagellated, rod-shaped bacteria of the Enterobacteriaceae family. The genus consists of three major species, Salmonella enterica (S. enterica), S. bongori, and S. subterranean. Salmonella in the gastrointestinal (GI) tract are pathogenic bacteria that trigger diarrhea, fever, vomiting, and abdominal cramps, and are sometimes related to postinfectious irritable bowel syndrome [41]. In addition, Salmonellosis is a risk factor of inflammatory bowel disease (IBD) and Clostridium difficile infection, which damages intestinal mucosal tissues [42,43]. S. enterica serotype Typhimurium (S. Typhimurium) triggers pro-inflammatory IL-8 expression and production via MAPK (in particular, p38) activation using the type III secretion system in IECs. The intake of a low concentration of DON renders IECs more susceptible to infection by S. Typhimurium and subsequent mucosal inflammatory responses owing to increased S. Typhimurium translocation [44,45,46]. Co-exposure to DON (≤0.5 μg/mL, in vitro) and Salmonella superinduces the expression of intestinal pro-inflammatory cytokines in porcine ileal tissues, resulting in detrimental inflammatory insults in humans and other animals [6]. Pigs exposed to a high dose of the type A trichothecene T-2 toxin via feed have decreased colonization of S. Typhimurium in the jejunum, ileum, and colon. However, a low concentration (1–100 ng/mL, in vitro) of T-2 toxin enhances the susceptibility of porcine macrophages and IECs to S. Typhimurium invasion [47]. Moreover, T-2 toxin-exposed pigs have increased translocation of S. Typhimurium through IEC monolayers. Although T-2 toxin favors infection by S. Typhimurium, it has adverse effects on the motility and metabolic activity of S. Typhimurium, suggesting both deleterious and favorable interactions between T-2 toxin and S. Typhimurium [48,49]. T-2 toxin has a profound negative effect on the ability of chickens to resist salmonellosis, but this is not accompanied by marked alterations in T- or B-cell responses to mitogenic stimulation [50]. In mice, increased mortality in response to S. Typhimurium challenge is dependent on T-2 toxin (≤1 mg/kg, in vivo) in a dose-dependent manner [51]. The increased mortality is mediated by increased splenic counts in mice challenged with S. Typhimurium, and T-2 toxin (1 mg/kg, in vivo) accelerates body-weight loss in infected mice [51]. In addition, challenge with T-2 toxin (1 mg/kg, in vivo) increases S. Typhimurium -related lesions in the spleens, kidneys, and livers, but Peyer’s patches and ileal tissues are marginally affected [52]. A high dose of OTA (3 mg/kg, in vivo) also increases intestinal colonization of S. Typhimurium in young chickens [53,54] and FB1 (150 mg/kg, in vivo) exacerbates clinical symptoms, such as diarrhea with bloody discharges, induced by S. enterica serotype Gallinarum (S. Gallinarum) infection [55]. Additionally, quail mortality is increased by FB1 (150 mg/kg, in vivo) from Fusarium moniliforme [55].
Macrophages play a crucial role in the pathogenesis of Salmonella infections because the bacteria are able to survive and multiply intracellularly after cellular entry. Macrophage invasion coincides with membrane ruffles, bacterium uptake, and the formation of Salmonella-containing vacuoles [56,57]. Although DON (0.25 μg/mL, in vitro) does not influence growth or the expression of S. Typhimurium virulence genes in macrophages, it promotes invasion and intracellular survival of S. Typhimurium in macrophages. Mechanistically, enhanced uptake of S. Typhimurium into macrophages by DON coincides with F-actin reorganization of cells. This is mediated by extracellular signal-regulated protein kinase 1/2 (ERK1/2), resulting in increased susceptibility of pigs to infection with S. Typhimurium [5]. Although peritoneal macrophages in mice exposed to T-2 toxin (0.1 μM, in vitro) do not influence lethality, lethality is significantly higher in S. Typhimurium-challenged mice exposed to T-2 toxin (2 mg/mL, in vitro) than in mice that are not exposed to T-2 toxin, suggesting that additional immune-related cells are involved in the pathogenesis of S. Typhimurium-induced lethality of hosts that are pre-exposed to T-2 toxin [58]. In most infection models, mycotoxins enhance Salmonella infections in macrophages, and increase inflammatory responses induced by Salmonella infections via the upregulation of pro-inflammatory cytokines and chemokines. In addition, Salmonella-induced inflammatory responses are enhanced by mycotoxins, and enhanced inflammatory responses may be a risk factor of inflammatory diseases, such as IBD. As a result of increased colonization, mycotoxins may promote Salmonella-induced pathogenicity, e.g., inflammatory responses, by synergistically inducing excessive pro-inflammatory cytokines or chemokine storms.

3.2. Relationship between Mycotoxin Exposure and Escherichia Coli Infection

Escherichia coli is a gram-negative, rod-shaped bacterium that is found in the lower intestine of mammals. Enterovirulent E. coli can be classified based on virulence and acquired genetic features. Enterotoxigenic E. coli (ETEC) produce one or more enterotoxins that are heat labile (LT-1 and LT-2) and secrete heat stable enterotoxins (STa and STb). Enteropathogenic E. coli (EPEC) harbor a pathogenicity island that encodes a series of proteins involved in the effacement lesion of the intestinal villi of cells. Enteroinvasive E. coli (EIEC) have biochemical, physiological, and generic properties that are similar to those of Sshigella, invading the epithelial cells of the colon. Enterohemorrhagic E. coli (EHEC) secrete Shiga-like toxins and cause severe diseases, such as bloody diarrhea and hemolytic uremic syndrome in humans [59,60,61,62,63,64,65,66,67,68,69,70]. A low level of aflatoxin (1–3 ppb, in vivo) or fumonisin (50–350 ppb, in vivo) increases cytotoxicity of Shiga-like toxin-producing E. coli in calves [71]. In pigs, dietary FB1 (0.5 mg/kg, in vivo) significantly increases colonization of EPEC strains in the small and large intestines, and subsequent bacterial translocation to extraintestinal organs, including the mesenteric lymph nodes, lung, liver, and spleen [72]. FB1 (1 mg/kg, in vivo) also downregulates antigen-specific immune responses induced by ETEC infections, resulting in longer shedding of ETEC following infection [73]. Combined exposure to FB1 (≤200 mg/kg, in vivo) and moniliformin (≤100 mg/kg, in vivo) (which is a mycotoxin produced by Fusarium species that regulates plant growth and is phytotoxic in maize and tobacco) reduces pathogenic E. coli clearance in poultry [74]. In broiler chickens, OTA (80 mg/kg, in vivo) worsens E. coli infection-mediated diseases, such as perihepatitis and pericarditis, and the fibrin layer becomes thicker after E. coli infection [75]. OTA feeding leads to severe swelling of the proximal convoluted tubules and degeneration of the tubular epithelium in E.coli-inoculated kidney. In this co-exposure model, degenerative and mononuclear cell infiltration are also observed in the liver [75]. Taken together, exposure to some mycotoxins interferes with host defenses against enterovirulent E. coli, leading to the failure of bacterial clearance and promoting mucosal colonization and invasion as well as inflammatory responses. Therefore, mycotoxin exposure can make hosts more susceptible to E. coli infection-mediated acute and chronic diseases, including hemolytic uremic syndrome and renal failure.

3.3. Relationship between Mycotoxin Exposure and Clostridium Perfringens Infection

Clostridium perfringens (C. perfringens) is a gram-positive, rod-shaped, spore-forming bacterium and is a risk factor for necrotic enteritis in broiler chickens [76,77,78,79]. C. perfringens has five toxinogenic types (A, B, C, D, and E) that are differentiated according to the production of four different toxins (Alpha, Beta, Epsilon, and Iota) [80]. Necrotic enteritis is one of the most important enteric diseases in poultry and is caused by type A C. perfringens isolates and rarely by type C isolates [81,82]. Type C toxins of C. perfringens are risk factors for several enteric diseases, such as hemorrhagic or necrotic enterotoxemia in piglets, lambs, and calves, and increase vaccination costs in the agricultural industry [81,83]. DON exposure (3000–4000 μg/kg, in vivo) at concentrations below the European maximum guidance level of 5000 μg/kg feed in poultry via food intake is a predisposing factor for severe intestinal barrier disruption and enhanced growth and toxin production of C. perfringens, resulting in the development of necrotic enteritis in broiler chickens [84]. Necrotic enteritis lesions are mainly distributed in the duodenum and jejunum; these are also the major absorption sites for DON, which affects the functions of the proximal part of the intestinal tract by reducing villus height in the duodenum. This is associated with impaired nutrient uptake or digestion via a reduction in differentiated IECs [84]. In addition, FB1 treatment (25 μg/mL, in vitro) enhances susceptibility to the toxicity of the epsilon toxin of C. perfringens, which mediates cell death in canine cells [85]. Taken together, severe enteric diseases caused by C. perfringens may be more likely after exposure to mycotoxins, particularly necrotic enteritis and hemorrhagic or necrotic enterotoxemia. Mechanistically, the increased predisposition to enteric disorders is associated with intestinal barrier disruption and increased cytotoxicity in the gut epithelium in response to mycotoxins.

3.4. Relationship between Mycotoxins and Reovirus Infection

Reoviruses (respiratory enteric orphan viruses) are nonenveloped dsRNA-containing viruses that are ubiquitous and infect many mammalian species, including humans and mice [86,87,88,89]. In humans, reoviruses are not associated with any diseases that are more severe than a mild enteric or respiratory illness; however, in mice and rats, they cause many disease syndromes affecting major vital organs, like the brain and heart [90,91]. Enteric reovirus infections produce a self-limited infection in which the virus is cleared from the GI tract within 7 to 14 days after onset [92,93]. Mucosal responses to enteric reovirus infections involve the induction of intestinal IgA and serum IgG as well as cellular immune responses [94,95,96,97,98]. The titers of infected reovirus in the GI tract are 10-fold higher in DON-exposed mice than in control mice after 5 days. In addition, DON (10 and 25 mg/kg, in vivo) increases the severity of the reovirus infection and shedding in feces as well as reovirus IgA responses. Elevated shifting to IgA occurs due to the suppression of Th1 and enhancement of Th2 cytokine expression in the presence of DON (10 and 25 mg/kg, in vivo). In addition, DON exposure (10 and 25 mg/kg, in vivo) makes viral clearance difficult in reovirus-infected hosts [99]. A high concentration of T-2 toxin (1.75 mg/kg, in vivo) also results in the suppression of reovirus-induced immune responses, and is associated with reduced clearance of the infected virus [100]. Thus, DON and T-2 toxin promote intestinal viral accumulation and the subsequent elevation of inflammatory insults to the host [100]. Trichothecene mycotoxins diminish cell-mediated viral clearance by enhancing IL-4, IL-6, and IL-10, despite the suppression of interferon-γ (IFN-γ) from Peyer’s patches [99,100]. Taken together, trichothecene mycotoxins are a predisposing factor for reovirus infections in the intestine owing to increases in innate and adaptive immunity, as evidenced by elevated pro-inflammatory cytokines, intestinal IgA, and serum IgG.

4. Interaction between Mycotoxins and Pathogen Infections in Respiratory Organs

Viral illnesses are the most common risk factors for upper respiratory symptoms, which affect the nasal cavity, paranasal sinuses, pharynx, and larynx [101]. In contrast, bacterial infections may develop after viral infections and affect upper and lower respiratory systems [101]. Bordetella bronchiseptica (B. bronchiseptica) and Pasteurella multocida (P. multocida) are highly prevalent airway pathogenic bacteria and contribute to multiple pathologies in respiratory diseases, such as pneumonia, which is an inflammatory condition of the lung that affects microscopic air sacs known as alveoli [102,103,104]. B. bronchiseptica infection prior to challenge with P. multocida results in colonization of the upper respiratory tract and tonsils. B. bronchiseptica-infected pigs exposed to FB1 (10 mg/kg, in vivo) show clinical signs including mild serous nasal discharge, sneezing, panting, and hoarseness, unlike pigs not exposed to FB1 [105]. P. multocida also increases the risk of pneumonia and the severity of the pathological changes [105]. Neither FB1 exposure (0.5 mg/kg, in vivo) nor infection with P. multocida affects weight gain or causes serious clinical signs or lung lesions, and both have minimal effects on the production of bronchoalveolar lavage fluid in pigs. However, co-treatment with FB1 and P. multocida leads to delayed growth, induced cough, increased bronchoalveolar lavage fluid-producing cells (such as macrophages and lymphocytes), and prominent lung lesions with increased levels of proinflammatory cytokines, such as TNF-α, IFN-γ, and IL-18 [106]. Another bacterium, Mycoplasma hyopneumoniae (M. hyopneumoniae), is a primary pathogen of enzootic pneumonia and causes huge economic losses in the pig industry [107]. FB1-fed (20 ppm, in vivo) pigs show enhanced infection with M. hyopneumoniae, which causes pulmonary inflammatory responses and subsequent severe illness that requires euthanasia and progressive pathology in pigs [108]. As another infective agent affected by fumonisin exposure, porcine reproductive and respiratory syndrome virus (PRRSV) is a pathogenic virus in pigs; it has an enveloped, positive-sense, and single-stranded RNA genome [109]. PRRSV is classified into genotypes 1 and 2 based on 3′-terminus structural genes or the entire genome [110,111]. Both genotypes cause reproductive failure, respiratory signs, and immunosuppression in pigs, and increase mortality of neonatal pigs [112]. PRRSV is an infectious virus that replicates within macrophages or monocytes with the lung. FB1 exposure (≤20 ppm, in vivo) aggravates the severe symptoms induced by PRRSV in a dose-dependent manner [113]. In conclusion, exposure of pigs to FB1 can increase the inflammatory responses to infection via the airway pathogens, affecting animal health and growth. Therefore, further studies are needed to investigate how these interactions between mycotoxins and pathogen infections affect host defense and disease progress to maximize agricultural production.

5. Effects of Mycotoxin-Pathogen Exposure on Chronic Mucosal Disorders

Chronic exposure to environmental mycotoxins exacerbates infectious pathogen-induced diseases. Some trichothecene mycotoxins, such as DON and T-2 toxin, can directly damage mucosal tissues via disruption of the gut epithelial barrier and subsequently facilitate the translocation of gut commensal microbiota, pathobionts, and pathogens. In particular, disrupted junctional proteins and epithelial cell death owing to mycotoxins may account for impaired epithelial barrier integrity, which in turn enhances bacterial or viral pathogen infections and subsequent inflammatory insults [114,115,116,117]. Moreover, mycotoxin exposure can be an etiological factor of environmental IBD and has similar mechanistic patterns, including disruption of the intestinal epithelial barrier due to the reduction and delocalization of junctional proteins, and facilitated luminal or external bacterial translocation, triggering excessive immune cell-related intestinal inflammation [44,45]. Airway exposure to mycotoxins may trigger pro-inflammatory responses observed in pneumonia and may facilitate infection of pneumonia-triggering pathogens, including B. bronchiseptica and P. multocida. In particular, P. multocida induces systemic infections and mediates cirrhosis, cellulitis, and increases risk level in patients with organ transplants. Moreover, chronic obstructive pulmonary disease found in the mucosa of larger bronchi is one of the most prevalent diseases in the world, and is exacerbated by bacterial infection [49]. Mycotoxin exposure is another etiological factor in the exacerbation of chronic obstructive pulmonary disease; it increases microbial infection, colonization, or excessive cytokine production via cytoskeletal reorganization mediated by various factors including myosin light chain kinase (MLCK) leading to pulmonary epithelial disruption, one of major pathological events in patients with chronic obstructive pulmonary disease (COPD) [118]. Although extensive studies have examined the association between mycotoxin exposure and infectious diseases, the specificity and chronicity of the related mucosal disorders still requires extensive observations with a broad spectrum of dose regimes and exposure durations in optimized animal models and animal-alternate systems.

6. Summary and Final Remarks on the Regulatory Implication

Mucosal exposure to mycotoxins in the GI and respiratory tracts has substantial effects on susceptibility to infectious agents. However, each mycotoxin leads to different biological events in response to mucosal infections, depending on the dose regime and host specificity (Table 1). (1) DON and T-2 toxin enhance susceptibility to S. Typhimurium infection by promoting its translocation and intercellular survival via F-actin reorganization in macrophages. In broiler chickens, DON enhances C. perfringens growth and toxin production. Reovirus-mediated immune responses are also regulated by exposure to DON in mice, including increased IgA responses, suppressed viral clearance, and Th2 cytokine polarization. (2) FB1 aggravates S. Gallinarum-induced bloody diarrhea and mortality in quails. In addition, FB1 reduces antigen-specific immune responses to pathogenic E. coli infections and its clearance. Moreover, FB1 exacerbates the severity of damages caused by respiratory tract-associated pathogens in pigs. (3) OTA also increases the colonization of S. Typhimurium in chickens, and aflatoxin increases EPEC/EHEC-mediated intoxication by enterotoxins and pore-forming toxins in calves.
Ultimately, studies on the interaction between mycotoxin and pathogens implicate the need for appropriate regulatory limits of mycotoxins in the food and feed for the human and animal population with underlying infectious diseases. Although the various levels of mycotoxin regulatory limits are already set up (Table 2), these are based on the toxicological risk assessment in a healthy population. People or animals with infectious diseases can be more resistant or susceptible to mycotoxicosis than the healthy population. Moreover, the mycotoxin-pathogen interaction in a population with non-pathogenic underlying diseases, including metabolic, cardiovascular, and oncological diseases, needs to be addressed in future investigations.
Table 1. Interactions between mycotoxins and mucosal pathogens.
Table 1. Interactions between mycotoxins and mucosal pathogens.
DONS. TyphimuriumIncrease in translocation [44,45,46]Pig
Increase in cytokine production [6]
Promotion of invasion and intracellular survival [47]Porcine
Increased susceptibility to infection by F-actin reorganization in macrophages [5]
T-2 toxinS. TyphimuriumEnhancement of infection [6]Pig
Reduction of bacteria mortality [48,49]
S. TyphimuriumIncrease in mortality [51]Mouse
Loss of body weight [51]
Increased lesions in spleen, kidney and liver [52]
S. TyphimuriumIncrease in colonization [53]Chicken
OTAS. TyphimuriumIncrease in colonization [53,54]Chicken
FB1S. GallinarumDiarrhea with bloody discharges and increase in mortality [55]Japanese quail
FB1EPEC/EHECIncrease enterotoxin and pore-forming toxin activity [71,73]Calf
EPECIncrease in colonization [72]Pig
Translocation to mesenteric lymph nodes, lung, liver, and spleen [72]
ETECReduction of antigen-specific immune responses via longer shedding of ETEC [73]Pig
FB1 and MoniliforminE.coliReduction of bacteria clearance [74]Poultry
AflatoxinEPEC/EHECIncrease in enterotoxin and pore-forming toxin activity [71]Calf
OTAE.coliSwollen proximal convoluted tubules, degradation of tubular epithelium, and intestinal mephritis in kidney [75]Broiler chicken
Degenerative and mononuclear cell infiltration in liver [71]
DONC. perfringensEnhancement of growth and toxin production [84]Broiler chicken
FB1C. perfringensEnhancement of susceptibility to epsilon-toxin [85]Canine
DONReovirusElevation of IgA responses [99]Mouse
Increase in severity of reovirus infection and shedding [99]
Suppression of Th1 cytokine expression [99]
Enhancement of Th2 cytokine expression [99]
Diminished cell-mediated viral clearance [99]
Elevation of reovirus-induced cytokine expression [99]
T-2 toxinReovirusSuppression of reovirus-induced immune responses [99]Mouse
Diminished cell-mediated viral clearance by suppression of IFN-γ and elevation of reovirus-induced cytokine expression [99,100]
FB1B. bronchiseptica and P. multocidaIncrease in extent and severity of pathogenic changes [102,103,104]Pig
B. bronchisepticaClinical changes including mild serous nasal discharge, sneezing, panting, and hoarseness [105]
P. multocidaDelayed growth, induced cough, and increased BALF cells such as macrophages and lymphocytes [106]
Increase in risk of pneumonia and severity of pathological changes [106]
Increase in TNF-α, IFN-γ and IL-18 expression [106]
M. hyopneumoniaeInduction of pulmonary inflammatory responses and enhancement of infection [108]
PRRSVSevere histological lesion of lung [113]
Table 2. Regulatory limits of food and feed mycotoxins in USA and EU.
Table 2. Regulatory limits of food and feed mycotoxins in USA and EU.
Foods or Feeds NationsMycotoxinProductsLimits (μg/kg)
FoodUSA (Action level)Aflatoxin (B1, B2, G1, G2)Brazil nut, peanut, peanut products, pistachio nut20
Aflatoxin M1Milk0.5
USA (Guidance level)Fumonisin (B1, B2, B3)Maize, maize products2000–4000
DeoxynivalenolWheat and wheat products1000
EUMycotoxinsCereal and its productsLimits (μg/kg)
B1B1 + B2 + G1 + G2M1
AflatoxinPeanut, nut products, grains, and maize2.0–12.04.0–15.0-
Raw milk and milk products--0.05
Grain products for infants and baby food0.1--
Products for infants--0.025
Ochratoxin AGrains, fruits, coffee beans, wine, and grape Juice2.0–10.0
Products for infants0.5
Fumonisin (B1+B2)Maize and its products200–4000
DeoxynivalenolNon-processed grain products1250–1750
Processed grain products500–750
Products for infants200
FeedUSAFumonisinAnimal feedHorses and rabbits1000
Cow, sheep and goat30,000
Ruminants and poultry15,000
Butchering poultry50,000
DeoxynivalenolGrains and grain products (>50% grain for cow or chicken)10,000
Grains and grain products (>40% grain for cow or chicken)5000
Grains and grain products digested by sheep5000
EUAflatoxin B1Complete feeding stuffs for cattle, sheep and goat 20
complete feeding stuffs for dairy animals5
complete feeding stuffs for calves and lambs10
Complete feeding stuffs for pigs and poultry (except young animals)20
Other complete feeding stuffs10
Complementary feeding stuffs for cattle, sheep and goats (except complementary feeding stuffs for dairy animals, calves and lambs)20
Complementary feeding stuffs for pigs and poultry (except young animals)20
Other complementary feeding stuffs5
Deoxynivalenolcereals and cereal products with the exception of maize by-products8000
maize by-products12,000
Complementary and complete feeding stuffs with the exception of maize by-products5000
complementary and complete feeding stuffs for pigs900
complementary and complete feeding stuffs for calves (<4 months), lambs and kids2000
Zearalenonecereals and cereal products with the exception of maize by-products2000
maize by-products3000
Complementary and complete feeding stuffs
complementary and complete feeding stuffs for piglets and gilts (young sows)100
complementary and complete feeding stuffs for sows and fattening pigs250
complementary and complete feeding stuffs for calves, dairy cattle, sheep (including lambs) and goats (including kids)500
Ochratoxin Acereals and cereal products 250
complementary and complete feeding stuffs for pigs50
complementary and complete feeding stuffs for poultry100
Fumonisin B1 and B2maize and maize products60,000
complementary and complete feeding stuffs for pigs, horses, rabbits and pet animals5000
complementary and complete feeding stuffs for fish10,000
complementary and complete feeding stuffs for poultry, calves (<4 months) and mink20,000
complementary and complete feeding stuffs for adult ruminants (>4 months) and mink50


This work was carried out with the support of the Cooperative Research Program for Agriculture Science & Technology Development (Project No. PJ01093206), Rural Development Administration, Republic of Korea.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Lee, J.; Kim, H.; Jeon, J.J.; Kim, H.S.; Zeller, K.A.; Carter, L.L.; Leslie, J.F.; Lee, Y.W. Population structure of and mycotoxin production by Fusarium graminearum from maize in South Korea. Appl. Environ. Microbiol. 2012, 78, 2161–2167. [Google Scholar] [CrossRef] [PubMed]
  2. Yang, G.H.; Jarvis, B.B.; Chung, Y.J.; Pestka, J.J. Apoptosis induction by the satratoxins and other trichothecene mycotoxins: Relationship to ERK, p38 MAPK, and SAPK/JNK activation. Toxicol. Appl. Pharmacol. 2000, 164, 149–160. [Google Scholar] [CrossRef] [PubMed]
  3. Sorenson, W.G.; Frazer, D.G.; Jarvis, B.B.; Simpson, J.; Robinson, V.A. Trichothecene mycotoxins in aerosolized conidia of Stachybotrys atra. Appl. Environ. Microbiol. 1987, 53, 1370–1375. [Google Scholar] [PubMed]
  4. Nusuetrong, P.; Pengsuparp, T.; Meksuriyen, D.; Tanitsu, M.; Kikuchi, H.; Mizugaki, M.; Shimazu, K.; Oshima, Y.; Nakahata, N.; Yoshida, M. Satratoxin H generates reactive oxygen species and lipid peroxides in PC12 cells. Biol. Pharm. Bull. 2008, 31, 1115–1120. [Google Scholar] [CrossRef] [PubMed]
  5. Vandenbroucke, V.; Croubels, S.; Verbrugghe, E.; Boyen, F.; de Backer, P.; Ducatelle, R.; Rychlik, I.; Haesebrouck, F.; Pasmans, F. The mycotoxin deoxynivalenol promotes uptake of Salmonella Typhimurium in porcine macrophages, associated with ERK1/2 induced cytoskeleton reorganization. Vet. Res. 2009, 40, 64. [Google Scholar] [CrossRef] [PubMed]
  6. Vandenbroucke, V.; Croubels, S.; Martel, A.; Verbrugghe, E.; Goossens, J.; van Deun, K.; Boyen, F.; Thompson, A.; Shearer, N.; de Backer, P.; et al. The mycotoxin deoxynivalenol potentiates intestinal inflammation by Salmonella typhimurium in porcine ileal loops. PLoS ONE 2011, 6. [Google Scholar] [CrossRef] [PubMed][Green Version]
  7. Rajagopalan, M.S.; Busch, M.P.; Blum, H.E.; Vyas, G.N. Interaction of aflatoxin and hepatitis B virus in the pathogenesis of hepatocellular carcinoma. Life Sci. 1986, 39, 1287–1290. [Google Scholar] [CrossRef]
  8. Wogan, G.N.; Kensler, T.W.; Groopman, J.D. Present and future directions of translational research on aflatoxin and hepatocellular carcinoma. A review. Food Addit. Contam. Part A Chem. Anal. Control Expo. Risk Assess. 2012, 29, 249–257. [Google Scholar] [CrossRef] [PubMed]
  9. Kimura, R.; Hayashi, Y.; Takeuchi, T.; Shimizu, M.; Iwata, M.; Tanahashi, J.; Ito, M. Pasteurella multocida septicemia caused by close contact with a domestic cat: Case report and literature review. J. Infect. Chemother. 2004, 10, 250–252. [Google Scholar] [CrossRef] [PubMed]
  10. Bondy, G.S.; Pestka, J.J. Immunomodulation by fungal toxins. J. Toxicol. Environ. Health B Crit. Rev. 2000, 3, 109–143. [Google Scholar] [PubMed]
  11. Moon, E.Y.; Rhee, D.K.; Pyo, S. In vitro suppressive effect of aflatoxin B1 on murine peritoneal macrophage functions. Toxicology 1999, 133, 171–179. [Google Scholar] [CrossRef]
  12. Jiang, Y.; Jolly, P.E.; Preko, P.; Wang, J.S.; Ellis, W.O.; Phillips, T.D.; Williams, J.H. Aflatoxin-related immune dysfunction in health and in human immunodeficiency virus disease. Clin. Dev. Immunol. 2008, 2008. [Google Scholar] [CrossRef] [PubMed]
  13. Boorman, G.A.; Hong, H.L.; Dieter, M.P.; Hayes, H.T.; Pohland, A.E.; Stack, M.; Luster, M.I. Myelotoxicity and macrophage alteration in mice exposed to ochratoxin A. Toxicol. Appl. Pharmacol. 1984, 72, 304–312. [Google Scholar] [CrossRef]
  14. Kozaczynski, W. Experimental ochratoxicosis A in chickens. Histopathological and histochemical study. Arch. Vet. Pol. 1994, 34, 205–219. [Google Scholar] [PubMed]
  15. Aleksandrov, M.; Dzhurov, A. Effect of ochratoxin on the health status of broilers. Vet. Med. Nauki 1987, 24, 38–43. [Google Scholar] [PubMed]
  16. Singh, G.S.; Chauhan, H.V.; Jha, G.J.; Singh, K.K. Immunosuppression due to chronic ochratoxicosis in broiler chicks. J. Comp. Pathol. 1990, 103, 399–410. [Google Scholar] [CrossRef]
  17. Ayed, I.A.; Dafalla, R.; Yagi, A.I.; Adam, S.E. Effect on ochratoxin A on Lohmann-type chicks. Vet. Hum. Toxicol. 1991, 33, 557–560. [Google Scholar] [PubMed]
  18. Thuvander, A.; Funseth, E.; Breitholtz-Emanuelsson, A.; Hallen, I.P.; Oskarsson, A. Effects of ochratoxin A on the rat immune system after perinatal exposure. Nat. Toxins 1996, 4, 141–147. [Google Scholar] [CrossRef] [PubMed]
  19. Harvey, R.B.; Elissalde, M.H.; Kubena, L.F.; Weaver, E.A.; Corrier, D.E.; Clement, B.A. Immunotoxicity of ochratoxin A to growing gilts. Am. J. Vet. Res. 1992, 53, 1966–1970. [Google Scholar] [PubMed]
  20. Lea, T.; Steien, K.; Stormer, F.C. Mechanism of ochratoxin A-induced immunosuppression. Mycopathologia 1989, 107, 153–159. [Google Scholar] [CrossRef] [PubMed]
  21. Kumar, M.; Dwivedi, P.; Sharma, A.K.; Sankar, M.; Patil, R.D.; Singh, N.D. Apoptosis and lipid peroxidation in ochratoxin A- and citrinin-induced nephrotoxicity in rabbits. Toxicol. Ind. Health 2014, 30, 90–98. [Google Scholar] [CrossRef] [PubMed]
  22. Sansing, G.A.; Lillehoj, E.B.; Detroy, R.W.; Miller, M.A. Synergistic toxic effects of citrinin, ochratoxin A and penicillic acid in mice. Toxicon 1976, 14, 213–220. [Google Scholar] [CrossRef]
  23. Neldon-Ortiz, D.L.; Qureshi, M.A. Effects of AFB1 embryonic exposure on chicken mononuclear phagocytic cell functions. Dev. Comp. Immunol. 1992, 16, 187–196. [Google Scholar] [CrossRef]
  24. Cusumano, V.; Rossano, F.; Merendino, R.A.; Arena, A.; Costa, G.B.; Mancuso, G.; Baroni, A.; Losi, E. Immunobiological activities of mould products: Functional impairment of human monocytes exposed to aflatoxin B1. Res. Microbiol. 1996, 147, 385–391. [Google Scholar] [CrossRef]
  25. Silvotti, L.; Petterino, C.; Bonomi, A.; Cabassi, E. Immunotoxicological effects on piglets of feeding sows diets containing aflatoxins. Vet. Rec. 1997, 141, 469–472. [Google Scholar] [CrossRef] [PubMed]
  26. Jakab, G.J.; Hmieleski, R.R.; Zarba, A.; Hemenway, D.R.; Groopman, J.D. Respiratory aflatoxicosis: Suppression of pulmonary and systemic host defenses in rats and mice. Toxicol. Appl. Pharmacol. 1994, 125, 198–205. [Google Scholar] [CrossRef] [PubMed]
  27. Osweiler, G.D.; Kehrli, M.E.; Stabel, J.R.; Thurston, J.R.; Ross, P.F.; Wilson, T.M. Effects of fumonisin-contaminated corn screenings on growth and health of feeder calves. J. Anim. Sci. 1993, 71, 459–466. [Google Scholar] [PubMed]
  28. Reddy, R.V.; Sharma, R.P. Effects of aflatoxin B1 on murine lymphocytic functions. Toxicology 1989, 54, 31–44. [Google Scholar] [CrossRef]
  29. Hamilton, P.B.; Harris, J.R. Interaction of aflatoxicosis with Candida albicans infections and other stresses in chickens. Poult. Sci. 1971, 50, 906–912. [Google Scholar] [CrossRef] [PubMed]
  30. Edds, G.T.; Nair, K.P.; Simpson, C.F. Effect of aflatoxin B 1 on resistance in poultry against cecal coccidiosis and Marek’s disease. Am. J. Vet. Res. 1973, 34, 819–826. [Google Scholar] [PubMed]
  31. Wyatt, R.D.; Ruff, M.D.; Page, R.K. Interaction of aflatoxin with Eimeria tenella infection and monensin in young broiler chickens. Avian Dis. 1975, 19, 730–740. [Google Scholar] [CrossRef] [PubMed]
  32. Berek, L.; Petri, I.B.; Mesterhazy, A.; Teren, J.; Molnar, J. Effects of mycotoxins on human immune functions in vitro. Toxicol. Vitr. 2001, 15, 25–30. [Google Scholar] [CrossRef]
  33. Marijanovic, D.R.; Holt, P.; Norred, W.P.; Bacon, C.W.; Voss, K.A.; Stancel, P.C.; Ragland, W.L. Immunosuppressive effects of Fusarium moniliforme corn cultures in chickens. Poult. Sci. 1991, 70, 1895–1901. [Google Scholar] [CrossRef] [PubMed]
  34. Forsell, J.H.; Witt, M.F.; Tai, J.H.; Jensen, R.; Pestka, J.J. Effects of 8-week exposure of the B6C3F1 mouse to dietary deoxynivalenol (vomitoxin) and zearalenone. Food Chem. Toxicol. 1986, 24, 213–219. [Google Scholar] [CrossRef]
  35. Robbana-Barnat, S.; Lafarge-Frayssinet, C.; Cohen, H.; Neish, G.A.; Frayssinet, C. Immunosuppressive properties of deoxynivalenol. Toxicology 1988, 48, 155–166. [Google Scholar] [CrossRef]
  36. Swamy, H.V.; Smith, T.K.; MacDonald, E.J.; Boermans, H.J.; Squires, E.J. Effects of feeding a blend of grains naturally contaminated with Fusarium mycotoxins on swine performance, brain regional neurochemistry, and serum chemistry and the efficacy of a polymeric glucomannan mycotoxin adsorbent. J. Anim. Sci. 2002, 80, 3257–3267. [Google Scholar] [PubMed]
  37. Harvey, R.B.; Kubena, L.F.; Huff, W.E.; Elissalde, M.H.; Phillips, T.D. Hematologic and immunologic toxicity of deoxynivalenol (DON)-contaminated diets to growing chickens. Bull. Environ. Contam. Toxicol. 1991, 46, 410–416. [Google Scholar] [CrossRef] [PubMed]
  38. Moon, Y.; Yang, H.; Lee, S.H. Modulation of early growth response gene 1 and interleukin-8 expression by ribotoxin deoxynivalenol (vomitoxin) via ERK1/2 in human epithelial intestine 407 cells. Biochem. Biophys. Res. Commun 2007, 362, 256–262. [Google Scholar] [CrossRef] [PubMed]
  39. Pinton, P.; Oswald, I.P. Effect of deoxynivalenol and other Type B trichothecenes on the intestine: A review. Toxins 2014, 6, 1615–1643. [Google Scholar] [CrossRef] [PubMed]
  40. Sugita-Konishi, Y.; Pestka, J.J. Differential upregulation of TNF-alpha, IL-6, and IL-8 production by deoxynivalenol (vomitoxin) and other 8-ketotrichothecenes in a human macrophage model. J. Toxicol. Environ. Health A 2001, 64, 619–636. [Google Scholar] [CrossRef] [PubMed]
  41. Schwille-Kiuntke, J.; Enck, P.; Zendler, C.; Krieg, M.; Polster, A.V.; Klosterhalfen, S.; Autenrieth, I.B.; Zipfel, S.; Frick, J.S. Postinfectious irritable bowel syndrome: Follow-up of a patient cohort of confirmed cases of bacterial infection with Salmonella or Campylobacter. Neurogastroenterol. Motil. 2011, 23, e479–e488. [Google Scholar] [CrossRef] [PubMed]
  42. Gradel, K.O.; Nielsen, H.L.; Schonheyder, H.C.; Ejlertsen, T.; Kristensen, B.; Nielsen, H. Increased short- and long-term risk of inflammatory bowel disease after Salmonella or Campylobacter gastroenteritis. Gastroenterology 2009, 137, 495–501. [Google Scholar] [CrossRef] [PubMed]
  43. Jess, T.; Simonsen, J.; Nielsen, N.M.; Jorgensen, K.T.; Bager, P.; Ethelberg, S.; Frisch, M. Enteric Salmonella or Campylobacter infections and the risk of inflammatory bowel disease. Gut 2011, 60, 318–324. [Google Scholar] [CrossRef] [PubMed]
  44. Sartor, R.B. Does Mycobacterium avium subspecies paratuberculosis cause Crohn’s disease? Gut 2005, 54, 896–898. [Google Scholar] [CrossRef] [PubMed]
  45. Sartor, R.B. Microbial influences in inflammatory bowel diseases. Gastroenterology 2008, 134, 577–594. [Google Scholar] [CrossRef] [PubMed]
  46. Goossens, J.; Pasmans, F.; Verbrugghe, E.; Vandenbroucke, V.; de Baere, S.; Meyer, E.; Haesebrouck, F.; de Backer, P.; Croubels, S. Porcine intestinal epithelial barrier disruption by the Fusarium mycotoxins deoxynivalenol and T-2 toxin promotes transepithelial passage of doxycycline and paromomycin. BMC Vet. Res. 2012, 8, 245. [Google Scholar] [CrossRef] [PubMed][Green Version]
  47. Verbrugghe, E.; Vandenbroucke, V.; Dhaenens, M.; Shearer, N.; Goossens, J.; de Saeger, S.; Eeckhout, M.; D’Herde, K.; Thompson, A.; Deforce, D.; et al. T-2 toxin induced Salmonella Typhimurium intoxication results in decreased Salmonella numbers in the cecum contents of pigs, despite marked effects on Salmonella-host cell interactions. Vet. Res. 2012, 43, 22. [Google Scholar] [CrossRef] [PubMed]
  48. Dimitroulia, E.; Pitiriga, V.C.; Piperaki, E.T.; Spanakis, N.E.; Tsakris, A. Inflammatory bowel disease exacerbation associated with Epstein-Barr virus infection. Dis. Colon Rectum 2013, 56, 322–327. [Google Scholar] [CrossRef] [PubMed]
  49. Erkan, L.; Uzun, O.; Findik, S.; Katar, D.; Sanic, A.; Atici, A.G. Role of bacteria in acute exacerbations of chronic obstructive pulmonary disease. Int. J. Chronic Obstr. Pulm. Dis. 2008, 3, 463–467. [Google Scholar]
  50. Ziprin, R.L.; Elissalde, M.H. Effect of T-2 toxin on resistance to systemic Salmonella typhimurium infection of newly hatched chickens. Am. J. Vet. Res. 1990, 51, 1869–1872. [Google Scholar] [PubMed]
  51. Tai, J.H.; Pestka, J.J. Impaired murine resistance to Salmonella typhimurium following oral exposure to the trichothecene T-2 toxin. Food Chem. Toxicol. 1988, 26, 691–698. [Google Scholar] [CrossRef]
  52. Tai, J.H.; Pestka, J.J. T-2 toxin impairment of murine response to Salmonella typhimurium: A histopathologic assessment. Mycopathologia 1990, 109, 149–155. [Google Scholar] [CrossRef] [PubMed]
  53. Fukata, T.; Sasai, K.; Baba, E.; Arakawa, A. Effect of ochratoxin A on Salmonella typhimurium-challenged layer chickens. Avian Dis. 1996, 40, 924–926. [Google Scholar] [CrossRef] [PubMed]
  54. Kubena, L.F.; Bailey, R.H.; Byrd, J.A.; Young, C.R.; Corrier, D.E.; Stanker, L.H.; Rottinghaust, G.E. Cecal volatile fatty acids and broiler chick susceptibility to Salmonella typhimurium colonization as affected by aflatoxins and T-2 toxin. Poult. Sci. 2001, 80, 411–417. [Google Scholar] [CrossRef] [PubMed]
  55. Deshmukh, S.; Asrani, R.K.; Jindal, N.; Ledoux, D.R.; Rottinghaus, G.E.; Sharma, M.; Singh, S.P. Effects of Fusarium moniliforme culture material containing known levels of fumonisin B1 on progress of Salmonella Gallinarum infection in Japanese quail: Clinical signs and hematologic studies. Avian Dis. 2005, 49, 274–280. [Google Scholar] [CrossRef] [PubMed]
  56. Finlay, B.B.; Ruschkowski, S.; Dedhar, S. Cytoskeletal rearrangements accompanying Salmonella entry into epithelial cells. J. Cell Sci. 1991, 99, 283–296. [Google Scholar] [PubMed]
  57. Monack, D.M.; Raupach, B.; Hromockyj, A.E.; Falkow, S. Salmonella typhimurium invasion induces apoptosis in infected macrophages. Proc. Natl. Acad. Sci. USA 1996, 93, 9833–9838. [Google Scholar] [CrossRef] [PubMed]
  58. Vidal, D.; Mavet, S. In vitro and in vivo toxicity of T-2 toxin, a Fusarium mycotoxin, to mouse peritoneal macrophages. Infect. Immun. 1989, 57, 2260–2264. [Google Scholar] [PubMed]
  59. Levine, M.M. Escherichia coli that cause diarrhea: Enterotoxigenic, enteropathogenic, enteroinvasive, enterohemorrhagic, and enteroadherent. J. Infect. Dis. 1987, 155, 377–389. [Google Scholar] [CrossRef] [PubMed]
  60. Jerse, A.E.; Yu, J.; Tall, B.D.; Kaper, J.B. A genetic locus of enteropathogenic Escherichia coli necessary for the production of attaching and effacing lesions on tissue culture cells. Proc. Natl. Acad. Sci. USA 1990, 87, 7839–7843. [Google Scholar] [CrossRef] [PubMed]
  61. Berlutti, F.; Casalino, M.; Zagaglia, C.; Fradiani, P.A.; Visca, P.; Nicoletti, M. Expression of the virulence plasmid-carried apyrase gene (apy) of enteroinvasive Escherichia coli and Shigella flexneri is under the control of H-NS and the VirF and VirB regulatory cascade. Infect. Immun. 1998, 66, 4957–4964. [Google Scholar] [PubMed]
  62. Nataro, J.P.; Kaper, J.B. Diarrheagenic Escherichia coli. Clin. Microbiol. Rev. 1998, 11, 142–201. [Google Scholar] [PubMed]
  63. Czeczulin, J.R.; Balepur, S.; Hicks, S.; Phillips, A.; Hall, R.; Kothary, M.H.; Navarro-Garcia, F.; Nataro, J.P. Aggregative adherence fimbria II, a second fimbrial antigen mediating aggregative adherence in enteroaggregative Escherichia coli. Infect. Immun. 1997, 65, 4135–4145. [Google Scholar] [PubMed]
  64. Boyce, T.G.; Swerdlow, D.L.; Griffin, P.M. Escherichia coli O157:H7 and the hemolytic-uremic syndrome. N. Engl. J. Med. 1995, 333, 364–368. [Google Scholar] [CrossRef] [PubMed]
  65. Bastian, S.N.; Carle, I.; Grimont, F. Comparison of 14 PCR systems for the detection and subtyping of stx genes in Shiga-toxin-producing Escherichia coli. Res. Microbiol. 1998, 149, 457–472. [Google Scholar] [CrossRef]
  66. Carter, A.O.; Borczyk, A.A.; Carlson, J.A.; Harvey, B.; Hockin, J.C.; Karmali, M.A.; Krishnan, C.; Korn, D.A.; Lior, H. A severe outbreak of Escherichia coli O157:H7—Associated hemorrhagic colitis in a nursing home. N. Engl. J. Med. 1987, 317, 1496–1500. [Google Scholar] [CrossRef] [PubMed]
  67. Karmali, M.A.; Petric, M.; Lim, C.; Fleming, P.C.; Arbus, G.S.; Lior, H. The association between idiopathic hemolytic uremic syndrome and infection by verotoxin-producing Escherichia coli. J. Infect. Dis. 1985, 151, 775–782. [Google Scholar] [CrossRef] [PubMed]
  68. Karmali, M.A.; Arbus, G.S.; Ish-Shalom, N.; Fleming, P.C.; Malkin, D.; Petric, M.; Cheung, R.; Louie, S.; Humphreys, G.R.; Strachan, M. A family outbreak of hemolytic-uremic syndrome associated with verotoxin-producing Escherichia coli serotype O157:H7. Pediatr. Nephrol. 1988, 2, 409–414. [Google Scholar] [CrossRef] [PubMed]
  69. Karch, H.; Tarr, P.I.; Bielaszewska, M. Enterohaemorrhagic Escherichia coli in human Enterohaemorrhagic Escherichia coli in human medicine. Int. J. Med. Microbiol. 2005, 295, 405–418. [Google Scholar] [CrossRef] [PubMed]
  70. Mellmann, A.; Bielaszewska, M.; Kock, R.; Friedrich, A.W.; Fruth, A.; Middendorf, B.; Harmsen, D.; Schmidt, M.A.; Karch, H. Analysis of collection of hemolytic uremic syndrome-associated enterohemorrhagic Escherichia coli. Emerg. Infect. Dis. 2008, 14, 1287–1290. [Google Scholar] [CrossRef] [PubMed]
  71. Baines, D.; Sumarah, M.; Kuldau, G.; Juba, J.; Mazza, A.; Masson, L. Aflatoxin, fumonisin and Shiga toxin-producing Escherichia coli infections in calves and the effectiveness of Celmanax(R)/Dairyman’s Choice applications to eliminate morbidity and mortality losses. Toxins 2013, 5, 1872–1895. [Google Scholar] [CrossRef] [PubMed]
  72. Oswald, I.P.; Desautels, C.; Laffitte, J.; Fournout, S.; Peres, S.Y.; Odin, M.; Le Bars, P.; Le Bars, J.; Fairbrother, J.M. Mycotoxin fumonisin B1 increases intestinal colonization by pathogenic Escherichia coli in pigs. Appl. Environ. Microbiol. 2003, 69, 5870–5874. [Google Scholar] [CrossRef] [PubMed]
  73. Devriendt, B.; Gallois, M.; Verdonck, F.; Wache, Y.; Bimczok, D.; Oswald, I.P.; Goddeeris, B.M.; Cox, E. The food contaminant fumonisin B(1) reduces the maturation of porcine CD11R1(+) intestinal antigen presenting cells and antigen-specific immune responses, leading to a prolonged intestinal ETEC infection. Vet. Res. 2009, 40, 40. [Google Scholar] [CrossRef] [PubMed]
  74. 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. [Google Scholar] [CrossRef] [PubMed]
  75. Kumar, A.; Jindal, N.; Shukla, C.L.; Asrani, R.K.; Ledoux, D.R.; Rottinghaus, G.E. Pathological changes in broiler chickens fed ochratoxin A and inoculated with Escherichia coli. Avian Pathol. 2004, 33, 413–417. [Google Scholar] [CrossRef] [PubMed]
  76. Barbara, A.J.; Trinh, H.T.; Glock, R.D.; Glenn Songer, J. Necrotic enteritis-producing strains of Clostridium perfringens displace non-necrotic enteritis strains from the gut of chicks. Vet. Microbiol. 2008, 126, 377–382. [Google Scholar] [CrossRef] [PubMed]
  77. Timbermont, L.; Haesebrouck, F.; Ducatelle, R.; van Immerseel, F. Necrotic enteritis in broilers: An updated review on the pathogenesis. Avian Pathol. 2011, 40, 341–347. [Google Scholar] [CrossRef] [PubMed]
  78. Baba, E.; Ikemoto, T.; Fukata, T.; Sasai, K.; Arakawa, A.; McDougald, L.R. Clostridial population and the intestinal lesions in chickens infected with Clostridium perfringens and Eimeria necatrix. Vet. Microbiol. 1997, 54, 301–308. [Google Scholar] [CrossRef]
  79. Parish, W.E. Necrotic enteritis in the fowl (Gallus gallus domesticus). I. Histopathology of the disease and isolation of a strain of Clostridium welchii. J. Comp. Pathol. 1961, 71, 377–393. [Google Scholar] [CrossRef]
  80. Fisher, D.J.; Fernandez-Miyakawa, M.E.; Sayeed, S.; Poon, R.; Adams, V.; Rood, J.I.; Uzal, F.A.; McClane, B.A. Dissecting the contributions of Clostridium perfringens type C toxins to lethality in the mouse intravenous injection model. Infect. Immun. 2006, 74, 5200–5210. [Google Scholar] [CrossRef] [PubMed]
  81. Songer, J.G. Clostridial enteric diseases of domestic animals. Clin. Microbiol. Rev. 1996, 9, 216–234. [Google Scholar] [PubMed]
  82. Engstrom, B.E.; Fermer, C.; Lindberg, A.; Saarinen, E.; Baverud, V.; Gunnarsson, A. Molecular typing of isolates of Clostridium perfringens from healthy and diseased poultry. Vet. Microbiol. 2003, 94, 225–235. [Google Scholar] [CrossRef]
  83. Walker, P.D.; Foster, W.H.; Knight, P.A.; Freestone, D.S.; Lawrence, G. Development, preparation and safety testing of a Clostridium welchii type C toxoid. I: Preliminary observations in man in Papua New Guinea. J. Biol. Stand. 1979, 7, 315–323. [Google Scholar] [CrossRef]
  84. Antonissen, G.; van Immerseel, F.; Pasmans, F.; Ducatelle, R.; Haesebrouck, F.; Timbermont, L.; Verlinden, M.; Janssens, G.P.; Eeckhaut, V.; Eeckhout, M.; et al. The mycotoxin deoxynivalenol predisposes for the development of Clostridium perfringens-induced necrotic enteritis in broiler chickens. PLoS ONE 2014, 9. [Google Scholar] [CrossRef] [PubMed][Green Version]
  85. Shimamoto, S.; Tamai, E.; Matsushita, O.; Minami, J.; Okabe, A.; Miyata, S. Changes in ganglioside content affect the binding of Clostridium perfringens epsilon-toxin to detergent-resistant membranes of Madin-Darby canine kidney cells. Microbiol. Immunol. 2005, 49, 245–253. [Google Scholar] [CrossRef] [PubMed]
  86. Bellum, S.C.; Dove, D.; Harley, R.A.; Greene, W.B.; Judson, M.A.; London, L.; London, S.D. Respiratory reovirus 1/L induction of intraluminal fibrosis. A model for the study of bronchiolitis obliterans organizing pneumonia. Am. J. Pathol. 1997, 150, 2243–2254. [Google Scholar] [PubMed]
  87. Chua, K.B.; Crameri, G.; Hyatt, A.; Yu, M.; Tompang, M.R.; Rosli, J.; McEachern, J.; Crameri, S.; Kumarasamy, V.; Eaton, B.T.; Wang, L.F. A previously unknown reovirus of bat origin is associated with an acute respiratory disease in humans. Proc. Natl. Acad. Sci. USA 2007, 104, 11424–11429. [Google Scholar] [CrossRef] [PubMed]
  88. London, L.; Majeski, E.I.; Paintlia, M.K.; Harley, R.A.; London, S.D. Respiratory reovirus 1/L induction of diffuse alveolar damage: A model of acute respiratory distress syndrome. Exp. Mol. Pathol. 2002, 72, 24–36. [Google Scholar] [CrossRef] [PubMed]
  89. Morin, M.J.; Warner, A.; Fields, B.N. Reovirus infection in rat lungs as a model to study the pathogenesis of viral pneumonia. J. Virol. 1996, 70, 541–548. [Google Scholar] [PubMed]
  90. Weiner, H.L.; Drayna, D.; Averill, D.R., Jr.; Fields, B.N. Molecular basis of reovirus virulence: Role of the S1 gene. Proc. Natl. Acad. Sci. USA 1977, 74, 5744–5748. [Google Scholar] [CrossRef] [PubMed]
  91. Sherry, B.; Schoen, F.J.; Wenske, E.; Fields, B.N. Derivation and characterization of an efficiently myocarditic reovirus variant. J. Virol. 1989, 63, 4840–4849. [Google Scholar] [PubMed]
  92. Barkon, M.L.; Haller, B.L.; Virgin, H.W.T. Circulating immunoglobulin G can play a critical role in clearance of intestinal reovirus infection. J. Virol. 1996, 70, 1109–1116. [Google Scholar] [PubMed]
  93. Rubin, D.H.; Kornstein, M.J.; Anderson, A.O. Reovirus serotype 1 intestinal infection: A novel replicative cycle with ileal disease. J. Virol. 1985, 53, 391–398. [Google Scholar] [PubMed]
  94. London, S.D.; Rubin, D.H.; Cebra, J.J. Gut mucosal immunization with reovirus serotype 1/L stimulates virus-specific cytotoxic T cell precursors as well as IgA memory cells in Peyer’s patches. J. Exp. Med. 1987, 165, 830–847. [Google Scholar] [CrossRef] [PubMed]
  95. Cuff, C.F.; Cebra, C.K.; Rubin, D.H.; Cebra, J.J. Developmental relationship between cytotoxic alpha/beta T cell receptor-positive intraepithelial lymphocytes and Peyer’s patch lymphocytes. Eur. J. Immunol. 1993, 23, 1333–1339. [Google Scholar] [CrossRef] [PubMed]
  96. Major, A.S.; Cuff, C.F. Effects of the route of infection on immunoglobulin G subclasses and specificity of the reovirus-specific humoral immune response. J. Virol. 1996, 70, 5968–5974. [Google Scholar] [PubMed]
  97. Silvey, K.J.; Hutchings, A.B.; Vajdy, M.; Petzke, M.M.; Neutra, M.R. Role of immunoglobulin A in protection against reovirus entry into Murine Peyer’s patches. J. Virol. 2001, 75, 10870–10879. [Google Scholar] [CrossRef] [PubMed]
  98. Virgin, H.W.T.; Bassel-Duby, R.; Fields, B.N.; Tyler, K.L. Antibody protects against lethal infection with the neurally spreading reovirus type 3 (Dearing). J. Virol. 1988, 62, 4594–4604. [Google Scholar] [PubMed]
  99. Li, M.; Cuff, C.F.; Pestka, J. Modulation of murine host response to enteric reovirus infection by the trichothecene deoxynivalenol. Toxicol. Sci. 2005, 87, 134–145. [Google Scholar] [CrossRef] [PubMed]
  100. Li, M.; Cuff, C.F.; Pestka, J.J. T-2 toxin impairment of enteric reovirus clearance in the mouse associated with suppressed immunoglobulin and IFN-gamma responses. Toxicol. Appl. Pharmacol. 2006, 214, 318–325. [Google Scholar] [CrossRef] [PubMed]
  101. Bosch, A.A.; Biesbroek, G.; Trzcinski, K.; Sanders, E.A.; Bogaert, D. Viral and bacterial interactions in the upper respiratory tract. PLoS Pathog. 2013, 9. [Google Scholar] [CrossRef] [PubMed]
  102. Brockmeier, S.L.; Palmer, M.V.; Bolin, S.R.; Rimler, R.B. Effects of intranasal inoculation with Bordetella bronchiseptica, porcine reproductive and respiratory syndrome virus, or a combination of both organisms on subsequent infection with Pasteurella multocida in pigs. Am. J. Vet. Res. 2001, 62, 521–525. [Google Scholar] [CrossRef] [PubMed]
  103. Galeziok, M.; Roberts, I.; Passalacqua, J.A. Bordetella bronchiseptica pneumonia in a man with acquired immunodeficiency syndrome: A case report. J. Med. Case Rep. 2009, 3, 76. [Google Scholar] [CrossRef] [PubMed]
  104. Klein, N.C.; Cunha, B.A. Pasteurella multocida pneumonia. Semin. Respir. Infect. 1997, 12, 54–56. [Google Scholar] [PubMed]
  105. Posa, R.; Donko, T.; Bogner, P.; Kovacs, M.; Repa, I.; Magyar, T. Interaction of Bordetella bronchiseptica, Pasteurella multocida, and fumonisin B1 in the porcine respiratory tract as studied by computed tomography. Can. J. Vet. Res. 2011, 75, 176–182. [Google Scholar] [PubMed]
  106. Halloy, D.J.; Gustin, P.G.; Bouhet, S.; Oswald, I.P. Oral exposure to culture material extract containing fumonisins predisposes swine to the development of pneumonitis caused by Pasteurella multocida. Toxicology 2005, 213, 34–44. [Google Scholar] [CrossRef] [PubMed]
  107. Maes, D.; Segales, J.; Meyns, T.; Sibila, M.; Pieters, M.; Haesebrouck, F. Control of Mycoplasma hyopneumoniae infections in pigs. Vet. Microbiol. 2008, 126, 297–309. [Google Scholar] [CrossRef] [PubMed]
  108. Posa, R.; Magyar, T.; Stoev, S.D.; Glavits, R.; Donko, T.; Repa, I.; Kovacs, M. Use of computed tomography and histopathologic review for lung lesions produced by the interaction between Mycoplasma hyopneumoniae and fumonisin mycotoxins in pigs. Vet. Pathol. 2013, 50, 971–979. [Google Scholar] [CrossRef] [PubMed]
  109. Bautista, E.M.; Faaberg, K.S.; Mickelson, D.; McGruder, E.D. Functional properties of the predicted helicase of porcine reproductive and respiratory syndrome virus. Virology 2002, 298, 258–270. [Google Scholar] [CrossRef] [PubMed]
  110. Murtaugh, M.P.; Elam, M.R.; Kakach, L.T. Comparison of the structural protein coding sequences of the VR-2332 and Lelystad virus strains of the PRRS virus. Arch. Virol. 1995, 140, 1451–1460. [Google Scholar] [CrossRef] [PubMed]
  111. Nelsen, C.J.; Murtaugh, M.P.; Faaberg, K.S. Porcine reproductive and respiratory syndrome virus comparison: Divergent evolution on two continents. J. Virol. 1999, 73, 270–280. [Google Scholar] [PubMed]
  112. Zimmerman, J.J.; Yoon, K.J.; Wills, R.W.; Swenson, S.L. General overview of PRRSV: A perspective from the United States. Vet. Microbiol. 1997, 55, 187–196. [Google Scholar] [CrossRef]
  113. Bane, D.P.; Neumann, E.J.; Hall, W.F.; Harlin, K.S.; Slife, R.L. Relationship between fumonisin contamination of feed and mystery swine disease. A case-control study. Mycopathologia 1992, 117, 121–124. [Google Scholar] [CrossRef] [PubMed]
  114. Diesing, A.K.; Nossol, C.; Danicke, S.; Walk, N.; Post, A.; Kahlert, S.; Rothkotter, H.J.; Kluess, J. Vulnerability of polarised intestinal porcine epithelial cells to mycotoxin deoxynivalenol depends on the route of application. PLoS ONE 2011, 6. [Google Scholar] [CrossRef] [PubMed]
  115. Diesing, A.K.; Nossol, C.; Panther, P.; Walk, N.; Post, A.; Kluess, J.; Kreutzmann, P.; Danicke, S.; Rothkotter, H.J.; Kahlert, S. Mycotoxin deoxynivalenol (DON) mediates biphasic cellular response in intestinal porcine epithelial cell lines IPEC-1 and IPEC-J2. Toxicol. Lett. 2011, 200, 8–18. [Google Scholar] [CrossRef] [PubMed]
  116. Klunker, L.R.; Kahlert, S.; Panther, P.; Diesing, A.K.; Reinhardt, N.; Brosig, B.; Kersten, S.; Danicke, S.; Rothkotter, H.J.; Kluess, J.W. Deoxynivalenol and E. coli lipopolysaccharide alter epithelial proliferation and spatial distribution of apical junction proteins along the small intestinal axis. J. Anim. Sci. 2013, 91, 276–285. [Google Scholar] [CrossRef] [PubMed]
  117. Pinton, P.; Tsybulskyy, D.; Lucioli, J.; Laffitte, J.; Callu, P.; Lyazhri, F.; Grosjean, F.; Bracarense, A.P.; Kolf-Clauw, M.; Oswald, I.P. Toxicity of deoxynivalenol and its acetylated derivatives on the intestine: Differential effects on morphology, barrier function, tight junction proteins, and mitogen-activated protein kinases. Toxicol. Sci. 2012, 130, 180–190. [Google Scholar] [CrossRef] [PubMed]
  118. Thorley, A.J.; Tetley, T.D. Pulmonary epithelium, cigarette smoke, and chronic obstructive pulmonary disease. Int. J. Chronic Obstr. Pulm. Dis. 2007, 2, 409–428. [Google Scholar]
Back to TopTop