* Department of Food Science and Human Nutrition,
Department of Microbiology,
Department of Veterinary Pathology,
§ Department of Pharmacology,
¶ Institute for Environmental Toxicology, and
|| National Food Safety and Toxicology Center, Michigan State University, East Lansing, Michigan 488241224
Received June 16, 1999; accepted October 20, 1999
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ABSTRACT |
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Key Words: trichothecene; vomitoxin; deoxynivalenol; mycotoxin; immunotoxicity; protein synthesis inhibition; spleen; endotoxin; flow cytometry; lipopolysaccharide; apoptosis; programmed cell death; thymus; Peyer's patch; bone marrow.
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INTRODUCTION |
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There is extensive evidence that LPS can influence the magnitude of toxic responses to xenobiotic agents. In some cases, LPS contamination of an environmental chemical source (e.g., LPS in machining fluids) may be the primary determinant of the toxic response (Gordon and Harkema, 1995; Mattsby-Baltzer et al., 1989
), whereas in other cases concurrent exposure to small amounts of LPS may magnify the inherent toxicity of a chemical. For example, exposure to modest, normally nontoxic doses of LPS markedly increases the hepatotoxic responses to a number of xenobiotic agents including CCl4, ethanol, galactosamine, and allyl alcohol (reviewed in Roth et al., 1997). These and other examples suggest that humans exposed to low doses of LPS may represent a subpopulation that is particularly sensitive to xenobiotic chemicals.The trichothecenes are a group of sesquiterpenoid fungal toxins that includes some of the most potent protein synthesis inhibitors known (Ueno, 1987
). Trichothecene mycotoxins are commonly found in cereal grains as a result of Fusarium infestation and have also been detected in air samples from water-damaged buildings that harbor the growth of Stachyobotrys (Johanning et al., 1996
). The trichothecene vomitoxin (VT or deoxynivalenol) is a common contaminant of wheat and corn products and can be found at ppm levels in ready-to-eat foods (Rotter et al., 1996
). Hallmarks of experimental or accidental high-dose trichothecene exposure include rapid diminution of lymphoid tissue and lymphopenia that precede death via a circulatory shocklike syndrome (Beardall and Miller, 1994
).
Apoptosis is a programmed mode of cell death that is essential in several biologic circumstances, including development of the immune system (Cohen et al., 1992). Cells undergoing apoptosis in vivo demonstrate nuclear and cytoplasmic condensation and dissolution into membrane-bound fragments that are phagocytosed by neighboring cells and rapidly degraded. A principal role for programmed cell death is thought to be efficient removal of stressed, damaged, or unnecessary cells from a tissue without the generation of inflammatory or immune responses. Exposure in vivo to large doses of LPS induces apoptosis in splenic germinal centers and thymus (THY) of mice (Zhang et al., 1994
) and swine (Norimatsu et al., 1995
). Recently, the induction of thymic atrophy with accompanying thymocyte apoptosis was reported in mice exposed to intraperitoneal injections of the trichothecenes T-2 toxin and fusarenone (Islam et al., 1998
).
In previous studies with mice, it has been observed that trichothecenes become more toxic in the presence of LPS, thereby causing markedly elevated tissue injury and mortality (Tai and Pestka, 1988; Taylor et al., 1991
; Zhou, et al., 1999
). Pronounced thymic and splenic lymphocyte depletion were characteristically observed in these studies. Because either LPS or trichothecenes alone will induce apoptosis in lymphoid tissue at large doses, we sought to evaluate the effects of low doses of Salmonella typhimurium LPS and a common food-borne trichothecene, VT, on apoptosis in four potential lymphoid targets, namely, thymus (THY), Peyer's patches (PP), spleen (SP), and bone marrow (BM). The study yielded both qualitative and quantitative evidence indicating that low doses of LPS and VT synergistically increase apoptotic cell loss in lymphoid tissue.
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MATERIALS AND METHODS |
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In a typical experiment, mice were given VT (25 mg/kg body weight) plus vehicle, LPS (0.5 mg/kg body weight) plus vehicle, VT (25 mg/kg body weight) plus LPS (0.5 mg/kg body weight), or the vehicles only. VT (Sigma, St. Louis, MO) was dissolved in 0.25 ml of tissue culture grade, endotoxin-free water (Sigma) and administered by a single oral gavage; animals that did not receive VT were treated with 0.25 ml water. LPS (Salmonella typhimurium, Sigma) was dissolved in 0.25 ml endotoxin-free water and was given by intraperitoneal injection; animals not treated with LPS were treated with 0.25 ml of endotoxin-free water. Twelve hours after exposure, mice were killed by cervical dislocation under ether anesthesia. THY, PP of intestine, SP, and femurs containing BM were immediately removed and processed for apoptosis measurements. The 25 mg/kg VT dose represents approximately one third of the LD50 for VT (Forsell et al., 1987). No mortality is observed at this dose (Zhou et al., 1999
).
DNA fragmentation assay.
Single cell suspensions were prepared according to the method of Islam et al. (1998). Following euthanasia by cervical dislocation, the THY, SP, PP, and femur were removed immediately from the mice. Single cells were released from THY, SP, and PP by mashing with a glass plunger against a fine stainless steel wire net (Collector Tissue Sieve, Bellco Glass Inc., Vineland, NJ) submerged into ice-cold PBS. BM was flushed out of the femur using PBS. The cells prepared from SP and BM were treated with erythrocyte-lysing buffer containing 0.83% (w/v) ammonium chloride, 0.1%(w/v) potassium bicarbonate, and 0.0037% (w/v) EDTA for 2 min at room temperature to remove erythrocytes. The cell suspension was passed through the 40-µm nylon sieve and the cell number was determined using a Bright-Line Hemacytometer (Sigma, St. Louis, MO).
DNA from THY, SP, PP, and BM was extracted and electrophoresed as described by Sellins and Cohen (1987). Briefly, cells (1 x 107) in phosphate-buffered saline (pH 7.4) were centrifuged for 5 min (500 x g) at 4°C, and the pellet was suspended in 0.1 ml hypotonic lysing buffer (10 mM Tris, 10 mM EDTA, 0.5% [v/v] Triton X-100, pH 8.0). Cells were incubated at 4°C for 10 min. The resultant lysate was centrifuged for 30 min (13,000 x g) at 4°C. Supernatant containing fragmented DNA was digested for 1 h at 37°C with RNase A (0.4 µg/µl). It was then incubated for 1 h at 37°C with proteinase K (0.4 µg/µl). DNA was precipitated in 50% isopropanol and 0.5 M NaCl overnight at 20°C. The precipitate was centrifuged at 13,000 x g for 30 min at 4°C. The resultant pellet was air dried, resuspended in 10 mM Tris, 1 mM EDTA, pH 8.0, then electrophoresed at 60 V for 2 h in 2% agarose gel (2 x 106 cells per lane) in 90 mM Tris-borate buffer (pH 8.0) containing 2 mM EDTA. After electrophoresis, the gel was stained with ethidium bromide (0.5 µg/ml), and the nucleic acids were visualized with a UV transilluminator. A 100-bp DNA ladder (GIBCO-BRL, Rockville, MD) was used as a molecular size marker.
Preparation of tissue sections.
For histologic assessment of apoptosis by cell morphology and detection of apoptotic cells in situ, tissues were fixed in 10% (v/v) buffered formalin for at least 24 h. Before embedding in paraffin, femurs were decalcified in 13% (v/v) formic acid for 3 days, and PP were pre-embedded in 3% (w/v) agarose followed by an additional 24 h fixation in 10% (v/v) buffered formalin. Tissues were embedded in paraffin and 4- to 6-µm sections were adhered to microscope slides. Sections were deparaffized and hydrated by heating the sections for 30 min at 60°C and transferring the slides through the following solutions: three times in xylene for 5 min, twice in 100% ethanol, and once in 90%, 80% ethanol, 5 min each, and then in distilled water.
Detection of apoptotic cells in situ.
A kit modification (Boehringer Mannheim in situ Cell Death Detection Kit, Indianapolis, IN) of the protocol of Gavrieli et al. (1992) for terminal deoxynucleotidyl transferase (TdT)-mediated fluorescein-dUTP nick-end labeling (TUNEL) was used to evaluate paraffin sections according to the manufacturer's instructions. Briefly, deparaffinized slides were rinsed with 10 mM phosphate-buffered saline (pH 7.0 PBS). TUNEL reaction mixture (50 µl) containing the components for the end-labeling reaction (fluorescein-dUTP and TdT) was added to each section previously encircled with a hydrophobic PAP Pen (Research Products International Corp., Mt. Prospect, IL). The section was overlaid with a coverslip to avoid evaporative loss, then incubated in a humidified chamber for 60 min at 37°C. After washing 3 times with PBS, samples were mounted with Gel/Mount (Biomedia Corp., Foster City, CA) and examined under a Nikon Labophot fluorescence microscope (Mager Scientific, Inc., Dexter, MI). Cells with nicked DNA were detectable by fluorescence. In each experiment, negative controls were included in which fixed and permeabilized samples were incubated within TUNEL reaction mixture devoid of TdT. Positive controls were also included that consisted of sections pretreated with DNase I (Sigma, St. Louis, MO), 1 mg/ml in buffer (30 mM Tris, pH 7.2, 140 mM sodium cacodylate, 4 mM MgCl2, 0.1 mM dithrothreitol), for 10 min at 37°C.
Histopathology and morphometry.
Paraffin sections (56 µm thick) of SP, THY, and PP were histochemically stained with hematoxylin/eosin (H&E) and examined by light microscopy (Olympus BX-60; Olympus Corp., Lake Success, NY). Upon histologic assessment of H&E-stained sections, cells were scored as apoptotic if they exhibited cellular shrinkage with concurrent cytoplasmic eosinophilia, nuclear pyknosis, and fragmentation (i.e., karyorrhexis) with associated apoptotic bodies. Digitized images of randomly selected fields (e.g., every 15th after a random start) were obtained (at a final magnification of 1550x for SP, 1580x for THY, and 950x for PP) and characterized morphometrically in terms of percent apoptotic lymphoid tissue. Images were captured and saved as PICT files on a Power Macintosh 7100/66 computer, using a high-resolution CCD camera (VE-1000CCD; Dage-MTI, Inc., Michigan City, IN) and the public domain image analysis program NIH Image v1.60 (written by Wayne Rasband, U.S. National Institutes of Health; available from the Internet by anonymous ftp from zippy.nimh.nih.gov). Randomly selected fields were captured so that approximately 5% of the total tissue section was analyzed for the THY (432 fields), 2.5% for SP (349 fields), and 50% for PP (237 fields). Only fields that were completely filled by tissue were counted (e.g., ones without large areas occupied by blood vessels, bordering spaces, etc.). The captured images were then imported to the morphometric data acquisition program Stereology Toolbox v1.1 (Morphometrix; Davis, CA) and analyzed using a double-density 25/125 point grid. The tissue directly beneath each grid point (THY and SP = 25 points/field, PP = 100 points/field) was categorized as normal lymphoid tissue, apoptotic lymphoid tissue, or "other" (e.g., nonlymphoid tissue, extracellular space, blood vessels, nonlymphoid cells, etc.). The number of points in each category was totaled individually for every animal. The area of each tissue category was estimated by the following calculation:
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Lymphoid tissue occupied approximately 6090% of the total tissue area examined.
Quantitation of apoptosis by flow cytometry.
Apoptosis in immune tissues was quantified by flow cytometric cell cycle analysis as described previously (Pestka et al., 1994). Briefly, THY, PP, SP, and BM cell suspensions were prepared as described for electrophoresis. Cells (2 x 106 )were resuspended in 0.2 ml PBS and 0.2 ml heat-inactivated fetal bovine serum, fixed by adding 1.2 ml ice-cold 70% (v/v) alcohol dropwise with gentle mixing and held overnight at 4°C. Cells were washed and incubated for 1 h in 1 ml propidium iodide (PI) DNA staining reagent (PBS, pH 7.4 containing PI 50 µg/ml, RNase 0.05 mg/ml at 50 units/mg, EDTA disodium 0.1 mM, and Triton x-100 0.1%) at room temperature and then stored on ice until analysis. The cell cycle distribution for single cells was measured with a Becton Dickinson FACS Vantage (San Jose, CA). Data from 10,000 cells were collected in list mode. The 488 line of an argon laser was used to excite PI; the fluorescence was detected at 620700nm. Cells were gated to exclude only debris and large cell aggregates and examined for DNA fluorescence intensity distribution. Cells in the DNA histogram with hypofluorescent DNA were designated apoptotic. All other cells distributed themselves in a normal cell cycle profile.
Statistics.
The data were analyzed using Sigma Stat for Windows (Jandel Scientific, San Rafael, CA). For comparisons of two groups of data, a Kruskal-Wallis One Way Analysis of Variance on Ranks and a Student-Newman-Keuls multiple comparison test was performed. Data sets showing significant differences (p < 0.05) were further analyzed for synergy. Single treatment replicates were randomly combined to calculate an expected mean additive response with variance. This calculated value was compared to actual cotreated samples using a Mann Whitney Rank Sum test.
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RESULTS |
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DISCUSSION |
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This paper is the first to show enhanced lymphocyte apoptosis resulting from combined trichothecene and LPS exposure. We chose to use the mouse model because its immune system is well characterized and we have extensively studied the immunologic effects of trichothecenes on this animal. Dose responses were tested in preliminary experiments using a small number of animals. The optimal doses for the observing synergy were 25 mg/kg VT and 0.5 mg/kg LPS. In preliminary experiments, we also examined 3, 6, 9, 12, and 24 h and obtained qualitative data to suggest that apoptosis was maximal at 12 h. To limit experimental animal use, we chose to focus on using on these two optimal doses at the 12-h timepoint to demonstrate the synergistic effect. The 25 mg/kg VT dose would represent the total daily dose that a mouse would acquire upon ingesting a diet contaminated with 100 ppm of this mycotoxin. Mice and rats are relatively resistant to LPS as compared to humans and other of animal species (Galanos et al., 1979). To study mechanisms of LPS toxicity in mice, various sensitization techniques such as galactosamime have been used (Redl et al., 1993
). When interpreted in this context, our data are valuable because they provide a basis for further mechanistic exploration of trichothecene-LPS interactions, dose response, and response kinetics, as well as assessment of this effect in other species using appropriate in vivo, ex vivo, and in vitro models. From such studies, the potential relevance to human health can be ascertained in the context of improved exposure data on the trichothecenes and LPS.
Systemic exposure of humans to LPS can occur through several mechanisms. The most widely recognized routes of LPS exposure are via respiratory and systemic Gram-negative bacterial infections (Brun-Buisson et al., 1995; Wenzel et al., 1996
). However, humans are also commonly exposed to the LPS of indigenous Gram-negative gut flora through gastrointestinal (GI) translocation, ie., the passage of LPS from the GI lumen into the blood (Jacob et al., 1977
). GI translocation of LPS is enhanced under a variety of conditions, including inflammatory bowel diseases (Palmer et al., 1980
), GI injury (Van Leeuwen et al., 1994
), liver disease (Bigatello et al., 1987
), dietary alterations (Spaeth et al., 1990
; Rutenburg et al., 1957
), and excessive alcohol consumption (Bode et al., 1987
). Elevated respiratory tract exposure to LPS also occurs in a variety of occupational environments. These include grain processing (Dosman et al., 1981
; Pernis et al., 1961
), waste treatment plants, machining operations (Mattsby-Baltzer et al., 1989
), poultry and swine industries (Donham et al., 1989
), and office or household air (Flaherty et al., 1984
; Peterson et al., 1964
; Rylander and Haglind, 1984
). Thus, exposure in humans is common, and the degree of exposure varies with occupation, diet, and disease state.
Exposure to large doses of LPS initiates a chain of inflammatory events that culminate in cell death, frank injury to tissues, and functional failure of several organs. LPS induces its marked biologic effects by stimulating host cells to produce a variety of mediators including proinflammatory cytokines (eg., TNF-, IL-6, IL-1), glucocorticoids, bioactive lipids, (eg., prostaglandins), reactive oxygen species, and activated coagulation components (Schletter et al., 1995
). Target cells for LPS are primarily mononuclear phagocytes but also include endothelial cells, neutrophils and smooth muscle cells. Of the aforementioned mediators, TNF-
appears to be of central importance (Beutler, 1995
). Exposure in vivo to large doses of LPS induces apoptosis in splenic germinal centers and thymus of mice and swine (Norimatsu et al., 1995
; Zhang et al., 1993
; 1994
). A key finding in these studies has been that elevated plasma levels of both TNF-
and glucocorticoids precede these effects, and lymphocyte apoptosis in mice can be blocked with neutralizing antibody to TNF-
. Accordingly, TNF-
appears to be an important factor in the genesis of apoptosis in lymphoid tissue during LPS exposure.
Superinduction is the capacity of protein synthesis inhibitors to augment and prolong the usually transient mitogenic induction of a gene as a secondary consequence of translational arrest. Superinduction is likely to involve transcriptional and/or post-transcriptional mechanisms (Li et al., 1997; Ouyang et al., 1996
). Trichothecenes inhibit translation (Ueno, 1987
), and we have demonstrated that VT superinduces mRNA expression and production of TNF-
and IL-6 in LPS-stimulated macrophages (Ji et al., 1998
; Wong et al., 1998
). Oral VT administration enhances TNF-
mRNA and protein expression in vivo within 2 h in spleen and other organs (Azcona-Olivera et al., 1995
; Zhou et al., 1997
), and this is potentiated by LPS coexposure (Zhou et al., 1999
). Thus, the enhanced lymphoid apoptosis observed herein may be mediated, in part, by elevated TNF-
level resulting from VT-mediated superinduction of LPS-induced TNF-
expression.
Our results are consistent with previous studies employing the trichothecene T-2 toxin. When LPS is administered at sublethal doses to mice, the estimated LD50 values for T-2 toxin markedly decrease (Tai and Pestka, 1988). Histologic analysis revealed that after coexposure of C3H/HeN mice to T-2 toxin (1 mg/kg) and LPS (2 mg/kg), extensive lymphocyte death occurred, whereas mice receiving only T-2 or LPS appeared normal. In related work, Taylor et al. (1991) subsequently observed synergy between T-2 toxin and LPS in mice. Effects included increased mortality, TNF-
production, hypothermia, and thymic atrophy. Plasma corticosterone concentration peaked at approximately 1 h for T-2 and LPS, with the LPS group being much greater (>2-fold); however, the combination group exhibited prolonged and elevated corticosterone concentration compared to LPS or T-2 alone. Taken together, the two studies suggest that T-2 toxin and possibly other trichothecenes become more toxic in the presence of LPS, thereby causing elevated tissue injury and mortality. The combination treatment appears to increase lymphoid organ depletion as well as increase two potential mediators of injury (TNF-
and corticosterone).
Flow cytometic analysis showed that there were greater percentages of hypofluorescent nuclei (indicative of apoptotic cells) in lymphocytes isolated from tissues of the combined VT/LPS treatment than in control or single-toxin treatment groups. Interestingly, although the morphometric and flow cytometric results yielded a similar outcome in this study, relative percentages of apoptotic cells estimated by flow cytometry differed from morphometric estimates in H&E sections. It is possible that flow cytometry may yield a more accurate estimate of apoptosis than morphometry because it samples the entire cell population of the lymphoid organ rather than sections. Using flow cytometry to measure the effects of VT and cycloheximide on lymphocyte apoptosis in vitro, Pestka et al. (1994) have observed that although both chemicals inhibit glucocorticoid-induced apoptosis in thymic and splenic T cells, these translational inhibitors induce apoptosis in B and IgA+ cells in SP and PP. These latter findings indicate that VT can either directly inhibit or enhance programmed cell death in a concentration-dependent manner and that its effect is highly dependent on lymphocyte subset, tissue source, glucocorticoid induction, and VT concentration. It would be of interest to study the phenotypic targets of the LPS/VT co-exposure model described here.
There are at least three general ways that LPS and VT may promote lymphocyte apoptosis. The first mechanism involves interaction between superinduced TNF- and lymphocytes. Direct engagement of TNF-
with its cell surface receptor has been shown to induce apoptosis (Baker and Reddy, 1996
; Hernandez-Caselles and Stutman, 1993
). A second mechanism involves TNF-
mediated elevation of soluble mediators. Likely candidates as secondary mediators include glucocorticoids, which induce apoptosis in both B and T cells (Pestka et al., 1994
; Garvy et al., 1993
), and prostaglandin E2 (PGE2) (Mohr et al., 1992
), which induces lymphocyte apoptosis (Brown and Phipps, 1996
). The third mechanism involves direct effects of VT on T and B cells, as has been suggested in our in vitro studies with VT (Pestka et al., 1994
). It should be emphasized that all three mechanisms are not mutually exclusive and could function simultaneously.
Persons exposed to LPS or other inflammagenic stimuli and who may thus be xenobiotic susceptible represent a significant part of the population (Roth et al., 1997). Because trichothecenes can be commonly found as food and indoor air contaminants, it is reasonable to suggest that a considerable portion of this sensitive human population may be exposed to these mycotoxins. Lymphocyte death induced by trichothecenes in LPS-exposed individuals might be expected to depress cell-mediated and humoral immune function. Characterization of phenotypic targets, dose-response, kinetics, strain/species sensitivity, and mechanisms in this model may provide insight into human health problems associated with trichothecene exposure in the xenobiotic-susceptible population.
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ACKNOWLEDGMENTS |
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NOTES |
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