72, 304-3 12 (1984)
and Macrophage Alteration Exposed to Ochratoxin A
G. A. BOORMAN, H. L. HONG, M. P. DIETER, H. T. HAYES, A. E. POHLAND,* M. STACK,* AND M. I. LUSTER National Toxicology Program, National Institute of Environmental Health Sciences, Research Triangle Park, North Carolina 27709, and *Bureau of Foods, Food and Drug Administration, Washington, D.C.
Received April 29, 1983; accepted August 15, 1983 Myelotoxicity
and Macrophage Alteration in Mice Exposed to Gchratoxin A.
G. A., HONG, H. L., DIETER, M. P., HAYES, H. T., POHLAND, M. I. (1984). Toxicol. Appl. Pharmacol. 72, 304-312. Six-
A. E., STACK, M., AND LUSTER,
to seven-week-old female B6C3Fr mice were administered a total of 0,20,40, or 80 mg/kg of ochratoxin A (OCT A) ip on alternate days over an S-day period. Twenty-four hours following the final dose, histopathology, bone marrow, and macrophage parameters were assayed.There was a dramatic dose related decrease in thymic mass with the mean thymus weight of the high dose animals being only 33% of controls. Histologic evidence of nephrotoxicity was minimal and restricted to the inner cortex. Myelotoxicity was present as evidenced by bone marrow hypocellularity, decreased marrow pluripotent stem cells (CFU-S), granulocyte-macrophage progenitors (CFIJ-GM& and decreased “Fe uptake in marrows and spleens of exposed mice. Peritoneal macrophages from sc as well as ip injected mice demonstrated increased phagocytic capacities and increased capacity to inhibit tumor cell growth. These alterations in bone marrow cells and macrophages suggest myelotoxicity is an additional potential hazard of GCT A exposure.
Ochratoxin A (OCT A) is a fungal toxin produced by Aspergillus ochruceus Wilhelm (van der Merwe et al., 1965) and Penicillium viridicatum Westling (van Walbeek et al., 1969). This mycotoxin occurs as a contaminant in wheat (Scott et al., 1970) and corn (Shotwell et al., 1969). Much of the previous work on ochratoxin focused on renal and hepatic toxicity (van Walbeek et al., 197 1; Purchase and Theron, 1968; Purchase and van der Watt, 1971; Doster et al., 1974; Suzuki et al., 1975; Krogh et al., 1979; Kanisawa et al., 1977; Szczech et al., 1973a,b; Hayes, 1981). Histopathologic changes consistently included renal tubular epithelial necrosis as well as hyaline degeneration and focal necrosis of the liver. Biochemical studies demonstrated that OCT A inhibits phosphorylase enzyme system apparently by competing with 3’,5’-cyclic AMP 0041~008X/84 $3.00 Copyright Q 1984 by Academic Press, Inc. All rights of reproduction in any form rexrwd.
for the enzyme phosphorylase b kinase (Pitout, 1968). The possible environmental hazard to humans has resulted in this compound being selected by the National Toxicology Program for long term testing in rodents. Following the OCT A exposure for 90 days (doses ranging from 0.6 to 1 mg/kg/day), both rats and mice showed histologic evidence of bone marrow hypocellularity (NTP data, unpublished). This observation was consistent with earlier studies where histologic evidence of bone marrow suppression and lymphoid depletion in spleens of ochratoxin-exposed chickens was found (Peckham et al., 1971). Several recent studies indicated that OCT A exposure in mice depressed humoral antibody responses (Prior and Sisodia, 1982; Creppy et al., 1983) suggesting functional changes in the immune system. The present report deals with histopathologic ob304
servations as well as bone marrow and macrophage functions in mice following subchronic OCT A exposure at dosages which did not induce overt nephrotoxicity. METHODS Mice. Female B6C3Fi (C57BL/6N X CsH) mice 6 to 8 weeks of age weighing 18 to 22 g were obtained through the National Cancer Institute production contracts (Charles River, Portage, Mich.). Animals were housed 10 per cage and allowed Freeaccessto commercial rodent chow (Ziegler Bros, Inc., Gardner, PA) and water. Ochratoxin A treatment. Ochmtoxin A (OCT A) was obtained from the National Cancer Institute (supplied by Food and Drug Administration). The sample was identified as OCT A by thin-layer and high performance liquid chromatography and quantified by gas chromatography as 92% pure. GCT A was freshly dissolved in 0.1 N sodium bicarbonate with 0.85% sodium chloride, protected from light, and stored in the refrigerator. Mice were injected ip or when indicated sc with OCT A in 0.2 ml volume on alternate days over an I-day period. The total doses administered were 20,40, or 80 mg/kg body weight. Control mice received isotonic 0.1 N sodium bicarbonate alone. Toxicological and immunological studies were performed 24 hr following the last treatment dose. Histopathology. Mice were killed with CO2 for necropsy. Body, liver, spleen, right kidney, and thymus weights were recorded, and tissues were collected for histopathology. Tissues were fixed in 10% buffered formalin, processed, and stained with hematoxylin and eosin. The lung, heart, liver, kidneys, adrenals, spleen, thymus, urinary bladder, skeletal muscle, bone marrow (sternum), and small and large intestines were examined histologically. Hematology. Blood samples were obtained via retroorbital venous plexus, and hematological parameters including hemoglobin, hematocrit, mean corpuscular volume, mean corpuscular hemoglobin, mean corpuscular hemoglobin concentration, red blood cell count (RBC), and total and differential white blood cell count (WBC) were examined. Cell numbers were determined in a Coulter particle counter, Model ZB (Coulter Electronics, Inc., Hialeah, Fl.). Clinical chemistry. Blood urea nitrogen (BUN), serum cholesterol, total bilirubin, total protein, albumin, serum glutamic pyruvic transaminase (SGPT), serum glutamic oxalic transaminase (SGOT), lactic acid dehydrogenase (LDH), and alkaline phosphatase were determined in a Centrifichem system P-1000 (Baker Instruments Corp., Pleasantville, N.Y.). Bone marrow cellularity. Marrow cells were aseptically collected by dissecting both femurs free of attached tissue, cutting the femur at the epiphysis, and flushing the shaft with 1.5 ml ofRPMI-1640 (GIBCO, Grand Island, N.Y.). Single cell suspensions were prepared by successive pas-
sagesof the cells through 22-gauge and 25-gauge needles. Nucleated cells were enumerated by the Coulter Model ZB Counter. Granulocyte-macrophage progenitors (CFU-GM). Progenitor cells (CFU-GM) horn the bone marrow and spleen were assayed by a modification of the method previously described by Bradley and Metcalf (1966). The culture medium consisted of RPME 1640 supplemented with 20Y0 fetal bovine serum, 5% human AB-type serum (Flow lab oratories, McLean. Va.), 2 mM L-glutamine (M. A. Bioproducts, Walkersville, Md.), 1% gentamycin, and 25 mM HEPES buffer (GIBCO, Grand Island, N.Y.). Nucleated femoral marrow cells (IO’) were suspended in 1.O ml of culture medium with 1.5% (w/v) methyl cellulose (Dow Chemical Co., Midland, Mich.) and 10% mouse lung conditioned medium (MLCM). Cells were added in quadruplicate to 35 X 10 mm petri dishes (Bioquest, Cockeysville, Md.). MLCM was prepared by culturing lungs from endotoxin-treated mice in serum free medium as previously mentioned (Sheridan et al.. 1975). CFUGM cultures were incubated for 7 days at 37°C in a humidified atmosphere containing 5 to 8% CO1 in air, and cells on the plates were then stained with methylene blue (Loefflers modified, Fisher Scientific Co., Raleigh, N.C.). The total number of colonies (240 cells) per plate were counted with a stcreomicroscope (Wild, Heerbrugg, Switzerland). A second measure of the bone marrow proliferative capacity was [3H]thymidine ([‘H]TdR) incorporation in liquid cultures. Bone marrow cells were added (1 X 105) to the culture medium in quadruplicate wells of a flat bottom microtiter plate (Linbro/Titertek, Flow Laboratories, McLean, Va.) and incubated for 4 days. The cells were then harvested by using a multiple cell harvester after a 4-hr pulse with [3H]TdR incorporation. The radioactivity retained on each dry filter was measured in 5 ml Hydroflour with a liquid scintillation counter (TRI-CARB 460 C, Packard Instrument Company, Downers Grove, Ill.). Colonyjbrming units in spleen (CFU-S). Pluripotent bone marrow stem cells were examined by the spleen colony method originally described by Till and McCulloch (1961). Isolated bone marrow cells (5 x 104) from OCI A exposed and control mice were injected iv into 2- to 3-month-old irradiated (600 rads) B6C3F, female recipients as previously described (Boorman et al., 1980). Recipients were killed 8 days following cell transfer; spleens were removed and fixed for 24 hr in Bouin’s fixative. Cell colonies were enumerated with a stereomicroscope. Histological sections were made of random colonies to confirm their myeloid nature. 59Fe uptake assay. 59Fe uptake was used to measure erythropoiesis by a modification of the method described by Boggs ez al. ( 1980). Control and OCT A-treated mice (seven/group) were injected ip with 0.5 &i of 5ve (specific activity 12.24 mCi/mg, New England Nuclear, Boston. Mass.) in 0.5 ml of 0.85% NaCl solution. Eighteen hours later the mice were killed with CO2 overdose, and radioactivity in the spleen and a whole leg from each mouse was determined in a Packard Autogamma Counter (PRIAS
Model BPGD, Packard Instrument Company, Downers Grove, Ill.). Bone marrow enzymology. Bone marrow was collected from paired femurs by flushing with sterile isotonic saline and single cell suspensions prepared by passage of the original solution through a 25-gauge needle. Cells were counted on a Coulter counter, and the solution was frozen at -65°C. After thawing, aliquots were sonicated in ice cold Tris buffer, pH 7.4, containing 0.1% T&on-X. Sonicated suspensions were centrifuged 5 min at 17,600g in 2.0-ml capped polypropylene tubes. Enzymes from the hexose monophosphate shunt (HMS) pathway, glycolysis or the Embden-Meyerhof (EM) pathway, and the tricarboxylic acid (TCA) cycle were assayed in bone marrow supematant fractions. Specific HMS enzymes were glucose6-phosphate dehydrogenase (EC 220.127.116.11, G6PD) and 6phosphogluconic dehydrogenase (EC 18.104.22.168, 6-PGD), EM enzymes were pyruvate kinase (EC 22.214.171.124, PK) and lactate dehydrogenase (EC 126.96.36.199, LDH), and TCA enzymes were isccitrate dehydrogenase (EC 188.8.131.52, ICDH) and malate dehydrogenase (EC 184.108.40.206, MDH). Assays were adjusted so that first order rates were attained. A centrifugal analyzer (Centrifichem, Union Carbide Co.) with an automated pipettor was used for the assays,and all reactions utilized coenzyme-linked standard methods (Bergmeyer, 1974) that measure the oxidation or reduction of NAD/NADP at 340 nm. Protein content in the supernate was measured by the Bradford-Coomassie blue dye-binding method (Bradford, 1976) with a commercial reagent (Bio-Rad Labs, Richmond, Calif.). Enzyme activities were related to protein concentration and are expressed as specific activity (nanomoles substrate converted/ minute/milligram protein). Macrophage assays. Macrophages were obtained from resident peritoneal cells following adherence to plastic microexudate llasks (Boorman ef al., 1980). The microculture growth inhibition assayfor murine macrophage activation was performed by a procedure previously described by Dean et al. (1978). Briefly, 20,000 MBL-2 leukemia target cells from 24-hr cultures were added in 0.1 ml volume to quadruplicate wells of round bottom 96-well plates, and macrophages from nontreated or GCI A-treated mice were added (0.1 ml) at macrophage to target cell ratios of 20: 1 and 10: 1. These cultures were incubated at 37°C in a humidified 5 to 7% CO* and air mixture for 48 hr. All cultures were washed, pulsed for 6 hr before termination with 1 &i of [3H]TdR (specific activity 6.7 Ci/ mmol, New England Nuclear, Boston, Mass.), and then the cells were harvested. The amount of incorporated [)H]TdR was determined in a Packard Model 460CD liquid scintillation counter. The percentage inhibition of target cell proliferation resulting from macrophage activation was calculated by the formula: 90 Growth Inhibition (% Cytostasis) cpm MBL-2 + macrophages from cpm MBL-2 + macrophages from control mice
ET AL. Phagocytosis was determined by examining uptake of fluorescent beads (Luster et al., 1982). Briefly, 0.4 X 106 resident peritoneal cells in 0.5 ml RPM1 culture medium were added to LabTek tissue culture chamber/slides (Miles Laboratories, Naperville, Ill.). The slides were incubated for 2 hr at 37’C, the medium was aspirated and replaced with 0.5 ml of fresh medium containing 4.5 X lo6 sonicated fluorescent beads (Covasphere Fx 1.4%-0.85 pm, red) (Covalent Technology Corp., San Jose, Calif.). The slides were incubated an additional 1 hr at 37°C on a rocker platform at low speed and washed thoroughly with medium; the cells were fixed in methanol. A minimum of 200 cells were read with a fluorescent microscope equipped with rhodamine lilters. Phagocytosis of three or more beads was considered positive. Statistical analysis. The student’s t test was employed to assessthe significance of treatment effectswhile doseresponse trends were determined by Jonckheere’s test (1954).
RESULTS Pathology OCT A exposure did not alter body, liver, kidney, or spleen weights as compared to controls, while the thymus showed a dose-related weight loss (Table 1). Histologically, the thymus showed marked cortical atrophy at the two highest doses with mild loss of cortical lymphocytes recognizable at the lowest dose level. Minimal histologic renal changes were present in all treatment groups with severity directly related to dosage. These lesions were restricted to the inner cortex and consisted of vacuolization and decreased staining intensity of renal tubular epithelial cells. In the two highest doses, the nuclei of a few renal tubular cells showed pyknosis and karyorrhexis. However, even these changes were mild and restricted to the inner cortex. An occasional mouse in the 80 mg/kg group showed acute toxicity and death within hours following the third or fourth dose of OCT A. There was no overt toxicity or mortality in the other dosage groups. Clinical
Clinical chemistry values are depicted in Table 2. Both blood urea nitrogen and serum
TABLE EFFECTS OFOCHRATOXINAON OCTA 0w.W
0 20 40 80
BUY weight k) 17.8 17.4 16.8 16.3
f + + T
0.5 0.6 0.4 0.6
33 67 23 42
53 55 53 57
2 2 t 2
I 3 2 I
62 74 72 69
+ 2 + f
(m.5) 2 + 2 +
3 2’ 3 3
949 946 892 933
3.5 4.3 4.3 4.3
k * t *
0.1 O.lb 0.2b 0.2”
Kidney (w) 121 128 116 131
t 8 _t 5 +-4 + 8
6.8 7.3 6.9 8.0
-+ + + _t
Thymus (mg) 0.3 0.4 0.2 0.3’
64 47 31 21
+ 5 ?I 6 +-2h t I”
Thymus/ MY 3.6 2.9 1.8 1.4
2 t 2 t
0.3 0.4 0.1” 0.1”
"X + SEM, seven mice per group. bp < 0.01 vs control. ‘p < 0.05 “S control.
alkaline phosphatase showed elevated results in the 40 and 80 mg/kg dosage groups. There was no effect on other serum chemistry values. Hematology
OCT A exposure had little significant biological effect on hematologic parameters (Table 3) on RBC indices (data not shown). The total counts of RBC and WBC remained unaltered; however, the number and percentage of neutrophils rose progressively while the percentage of lymphocytes was reduced significantly at the higher exposure. Hematopoiesis
The effect of OCT A on hematopoietic progenitor cells and erythropoiesis is shown in Tables 4 and 5. There was a significant (p < 0.01) dose-related treatment effect with reTABLE
spect to the reduction of bone marrow macrophage-granulocyte progenitors. Bone marrow cellularity also exhibited a significant (p < 0.01) dose-dependent decrease in both the 40 and 80 mg/kg exposure groups. The number of hematopoietic pluripotent stem cells (CFU-Ss) demonstrated a significant (p < 0.01) reduction in all OCT A-treated groups compared to the controls. The [3H]TdR incorporation in liquid bone marrow cultures, a measure of CFU-GM kinetics, was also depressed at the higher OCT A dosage levels. Erythropoiesis, as measured by 59Fe uptake, was also reduced in the femur at all three dosage levels, with the 80 mg/kg group being 56% less than controls (Table 5). “Fe uptake in the spleen was depressed in the highest dose group while an apparent compensatory increase occurred in the lower dosage groups being enhanced in both 20 and 40 mg/kg groups. There were significant dose-related increases in hexose monophosphate shunt enzyme activities from bone marrow of ochratoxin Atreated mice. Maximal increases plateaued in the 40 mg/kg treatment group at levels twofold above controls (Table 6). Macrophage
0 20 40 80
26.8 + I.1 29.5 f 3.1 39.1 + 4.2’ 40.8 + l.lb
’ x‘ f SEM, six mice/group. bp < 0.01 vs control.
97? 90% 102 AI 142 +
12 10 12 21
32 27 26 31
+ 3 2 2 + 2 +-2
77f 4 70* 7 92* 8 158 f 14b
The ability of OCT A to modulate macrophage activity was determined by examining total peritoneal cell numbers as well as phagocytic and tumor cell cytostatic activity of adherent resident peritoneal cells (>95% mac-
Differential OCTA (mg/kg) 0 20 40 80
RBCs X (10m6/mm3) 8.2 7.6 8.0 8.1
+ + + f
0.2” 0.3 0.1 0.2
7.2 4.3 1.8 8.2
+ + + k
1.6 0.5 1.4 1.9
6.0 3.5 4.7 3.2
k -t f +
1.3 0.5 0.6 0.8
Neutrophils (X 10m3/mm3
0.7 0.6 2.5 4.3
(84%) (81%) (62%)* (43%)’
+ zk + +
0.2 0.2 0.6’ 1.3’
(10%) (16%) (31%)b (509)b
Monocytes (W only)
Eosinophils (% only)
(1%) (3%) (3%)
a X f SEM, seven mice/group. * p -z 0.0 1 vs control. cp < 0.05 vs control.
rophages as determined by nonspecific esterase staining). The data presented in Table 7 represent values obtained in mice administered OCT A through SCinjection although similar results for phagocytosis and tumor cell cytostasis were obtained following ip administration. Resident peritoneal cell numbers were slightly but not significantly increased in treated mice. However, OCT A treatment induced the appearance of inflammatory macrophages as evidenced by increased phagocytic capacity and cytostatic activity at the 40 and 80 mg/kg dose level.
for myelotoxicity and immunotoxicity is less well defined (Peckham et al., 197 1). Recently evidence has been presented for altered B lymphocyte function (Creppy et al., 1983). Since overt toxicity may cause secondary alterations in bone marrow and immune function, this study was designed to evaluate ochratoxin exposure levels not causing severe parenchymal organ damage. Only minimal evidence of nephrotoxicity was found by histopathology and clinical chemistry. However some mortality was observed in the highest exposure group. Two animals dying within hours of the first dose showed no specific morphological alterations. A third animal dying after three doses was markedly dehydrated and had mild tubular necrosis. Altered bone marrow progenitor cells and decreased marrow cellularity were found at the 20 and 40 mg/ kg dosage levels.
DISCUSSION While the adverse effect of ochratoxin on the liver, kidney, and fetus is well known (Szczech et al., 1973b; Hayes, 1981), evidence TABLE EFFECT OF OCHRATOXIN OCT A @x/W 0 20 40 80
Nucleated cells/ femur (X10m6) 21.1 21.8 17.6 14.8
+ k k f
0.5 1.1 0.5* 1.0*
’ X k SEM, seven mice/group. * p < 0.01 vs control. L p < 0.05 vs control.
A ON BONE MARROW CFU-GM/ marrow 74 f 66 * 57+ 50*
10’ cells 1 2* lh I*
CF’U-GM/femur (X lo-*) 156 142 99 74
+ k +f
AND HEMOPOIETIC CFU-s/5 x lo4 cells injected
5 5 3h 5*
17.0 13.6 10.9 10.5
* k * *
0.2 0.3h 0.3* 0.4h
PROGENITORS” [‘H]TdR cpm/2 9185 8136 7195 7677
incorporation X IO4 cells f f k -t
321 579 484h 379’
TABLE 5 Emcr
A ON ERYTHROPOIESIS~
59Fe incorporation OCT A @g/kg) 0 20 40 80
Bone marrow 2479 1854 1342 1084
f + + f
229 82 100b 116’
1419 1818 2890 828
rf: 297 2 203 k 220b + 74
a X cpm k SEM, seven mice/group. bp < 0.01 vs control.
The present study indicates that OCT A exposure caused decreased pluripotent stem cells and granulocyte-macrophage progenitors, and affected the erythroid series as well, since decreased 59Fe uptake in bone marrow and spleen of exposed mice occurred. An interesting result in the 59Fe assay was the enhanced uptake in the spleen at lower OCT A doses which would suggest that splenic erythropoiesis is compensating for bone marrow decline in erythropoiesis. This compensation has been reported by others following exposure to myelotoxic agents (Silini et al., 1976). OCT A induced myelotoxicity may reflect a subtle indicator of general toxicity rather than a specific effect on bone marrow cells. OCT A is an effective inhibitor of protein synthesis presumably via inhibition of phenylalanyl-tRNA synthetase (Creppy et al., 1979, 1980; Haubeck et al., 1981) and thus likely to inhibit rapidly proliferating ceils such as bone marrow progenitors. In this study we demonstrated that
alterations in bone marrow progenitors occurs at exposure levels where only minimal parenchymal organ toxicity is seen. Cells of the bone marrow constitute a multipotential stem cell pool at various stages of differentiation, along with mature erythrocytes, thrombocytes, and leukocytes. Here we have examined enzymes from several pathways of glucose metabolism because that substrate constitutes a major energy source for both leukocytes (Jemelin and Frei, 1970; Lengle et al., 1978) and erythrocytes (DeBruin, 1976) and because the HMS pathway in bone marrow is particularly sensitive to other immunotoxicants like mercuric chloride (Dieter et al., 1983) and diethylstilbestrol (DES) (Luster et al., 1983; Dieter and French, 1983). Recent studies with DES (Luster et al., 1983) resulted in decreased bone marrow cellularity and progenitor cell activity, but in sharp contrast to the present study, those changes were closely correlated with a decrease in bone marrow HMS metabolism. In an extension of the DES study, we were able to separate the bone marrow into more discrete cell populations and found that depressed HMS metabolism by DES was wholly a reflection of enzyme inhibition in granulocytes and not in lymphocytes or erythrocytes (Dieter and French, 1983). Ochratoxin A inhibits protein synthesis (Haubeck et al., 1981) and thus might be expected to inhibit HMS enzyme activities in bone marrow, since the function of this pathway is to provide NADPH for reductive biosyntheses (Horecker and Hiatt, 1958) and
TABLE 6 EFFECT OF OCHRATOXIN
OCTA O-Wk) 0 20 40 80
G6PD 29 f 4Ok 53 f. 51 +
I lb 4b 36
6PGD 8.9 12.5 16.6 15.7
‘X f SEM, Seven mice/group. bp i 0.0 1 vs control.
5~ 0.3 f 0.76 f I.46 + 0.9b
A ON BONE MARROW
PK (mmol/min/mg 11.9 + 1.4 12.4 IL 1.4 11.6 + 3.3 8.3 k 0.7
LDH protein) 312 369 345 314
+ k 2 f
21 23 50 II
ICDH 12.6 15.2 15.3 14.8
f 1.3 + 1.5 +- 1.8 f 1.4
MDH 737 809 645 648
+ 34 f 63 2 28
TABLE 7 EFFECT OF
A ON RESIDENT
No. resident peritoneal cells (x10-6)
Macrophage phagocytosis (%I
173 + 4= (33%1)
MBL-2 cytostasis/ cpm + SEM (X 10e3) (% cytostasis) + 2
I ?O' (99%T)
a Each value represents a pool of IO mice/dosage group. b Significantly different from control at p < 0.05. ’ Significantly different from control at p < 0.01.
pentose sugars for incorporation into nucleic acids (Burt and Wenger, 196 1). Here the lack of correlation between depressed bone marrow cellularity and HMS metabolism suggests an entirely different explanation for the HMS enzyme responses to acute ochratoxin A treatment. Present results could be because of increased specific enzyme activity per cell, but more likely reflect enhanced HMS metabolism from increased numbers of an entirely different bone marrow cell population, perhaps osteoblasts or reticulocytes. Lead toxicity, for example, increased the proportion of circulating reticulocytes which resulted in an increase in blood glucose-6-phosphate dehydrogenase enzyme activity (DeBruin, 1976). The heterogeneity of the bone marrow cell populations assayed here does not allow us to determine which cell populations were affected, but the magnitude and dose-response to ochratoxin A support the validity of the biological response. The elevated activities may also be a rebound phenomenon after initial depression since the enzyme assays were conducted 24 hr after the last of four 24-hr staggered injections of OCT A. Techniques for sorting bone marrow cells into individual populations will eventually allow more precise analysis of enzyme pathway alterations.
We observed a dose-response increase in macrophage activation, i.e., increased phagocytosis and tumor cell cytostasis following OCT A treatment. It is generally accepted that mycotoxins are more toxic to the reticuloendothelial system than to lymphocytes (Richard et al., 1978). However, since macrophages serve as accessory cells for most immune functions, perturbations in macrophage activity may ultimately be reflected as altered cellular or humoral immune functions. This outcome appears to be the case in mice treated with the mycotoxin, Fusarenon-x, where treated mice possess high levels of regulatory suppressor macrophages which depress specific immunity (Masuda et al., 1982). In this respect, recent studies have indicated that OCT A-treated mice have decreased antibody response (Prior and Sisodia, 1982; Creppy et al., 1983). Studies in our laboratory have provided evidence that OCT A depresses antibody responses only at dose levels that approach levels which induce overt toxicity (unpublished data). The mechanisms associated with OCT A-induced macrophage peturbations are unknown but may be indirect effects resulting from altered myelogenous development particularly with respect to CFU-GM kinetics or due to inhibition of protein synthesis via in-
hibition of phenylalanyl-tRNA synthetase (Hayes et al., 1977; Creppy et al., 1979). This study demonstrated that the bone marrow cells as well as macrophages obtained from OCT A-treated mice were functionally altered and that these changes are found at dose levels that cause only mild toxicity in other organ systems. It would appear that this common mycotoxin has the potential to alter immunity and bone marrow function.
DeBruin, ed.). pp. 1289- 1297. Elsevier-North Holland, New York. DIETER, M. P., AND FRENCH, J. E. (1983). Estrogenic inhibition of carbohydrate metabolism in mouse bone marrow. J. Cell B&hem. (Suppl. 7B), 22. DIETER, M. P., LUSTER, M. I., BOORMAN, G. A., JAMESON, C. W., DEAN, J. H., AND Cow, J. W. (1983). Immunological and biochemical responses in mice treated with mercuric chloride. Toxicol. Appl. Pharmacol. 68, 218-228. DOSTER, R. C., SINNHUBER, R. O., AND PAWLOWSKI, N. E. (1974). Acute intraperitoneal toxicity of ochratoxin A and B derivatives in rainbow trout. Food Cosmet. Toxicol. 12, 499-505. REFERENCES HAUBECK, H. D., LQRKOWSKI, G., KOLSCH, E., AND R~SCHENTHALER, R. (198 1). Immunosuppression of BERGMEYER, H. U. (1974). Methods of Enzymatic Analochmtoxin A and its prevention by phenylalanine. Appl. ysis, 2nd Ed. Academic Press, New York. Environ. Microbial. 41, 1040- 1042. BRADFORD, M. M. (1976). A rapid and sensitive method HAYES, A. W. (198 1). Mycotoxin Teratogenicity and Mufor the quantitation of microgram quantities of protein tagenicity. CRC Press, Boca Raton, Florida. utilizing the principle of protein-dye binding. Anal. HAYES, A. W., CAIN, J. A., AND MOORE, B. G. (1977). B&hem. 72, 248-254. Effect of aflatoxin B,, ochratoxin A and rubratoxin B BRADLEY, T. R., AND METCALF, D. (1966). The growth on infant rats. Food Cosmet. Toxicol. 15, 23-27. of mouse bone marrow in vitro. Aust. J. Exp. Biol. Med. HORECKER, B. L., AND HIA=, H. H. (1958). Pathways Sci. 44,287-300. of carbohydrate metabolism in normal and neoplastic Bcxzi~s, D. R., BOGGS, S. S., CHERVENICK, P. A., AND cells. New Engl. .I Med. 258, 177-I 84. PATRENE,K. D. (1980). Murine recovery from busulfanJEMELIN, M., AND FREI, J. (1970). Leukocyte energy meinduced hematopoietic toxicity as assessedby three astabolism. III. Anaerobic and aerobic ATP production says for colony-forming cells. Amer. J. Hemat. 8, 43. and related enzymes. Enzyme Biol. Clin. 11, 289-323. BOORMAN, G. A., LUSTER, M. I., DEAN, J. H., AND WILSON, R. E. (1980). The effect of adult exposure to di- JONCKHEERE,A. R. (1954). A distribution-free K-sample test against ordered alternatives. Biometrika 41, 133ethylstilbestrol in the mouse: Macrophage function and 145. numbers. J. Reticuloendothel. Sot. 28, 547-560. KANISAWA, M., SUZUKI, S., KOZUKA, Y., AND YABURT, A. M., AND WENGER, B. S. (1961). GlucosedMAZAKl, M. (1977). Histopathological studies of the phosphate dehydrogenase activity in the brain of the toxicity of ochratoxin A in rats. 1. Acute oral toxicity. developing chick. Develop. Biol. 3, 84-95. Toxicol. Appl. Pharmacol. 42, 55-64. CREPPY,E. E., LUGNIER, A. A. J., FASIOLO, F., KELLER, KROGH, P., ELLING, F., FRIIS, C., HALD, B., LARSON, H., R~SCHENTHALER, R., AND DIRHEIMER, G. (1979). A. E., LILLE, H. J., MADSEN, A., MORTENSEN, P., RASIn vitro inhibition of yeast phenylalanyl-tRNA syntheMUSSEN,F., AND RAUNSKOV, U. (1979). Porcine netase by ochratoxin A. Chem. Biol. Interact. 24, 257phropathy induced by long-term ingestion of ochmtoxin 261. A. Vet. Pathol. 16, 466-475. CREPPY,E. E., SCHLEGEL,M., R~SCHENTHALER, R., AND LENGLE, E. E., GU~TIN, N. C., GONZALEZ, F., MENAHAN, DIRHEIMER, G. ( 1980). Phenylalanine prevents acute L. A., AND KEMP, R. G. (1978). Energy metabolism poisoning by ochratoxin A in mice. Toxicol. Left. 6, in thymic lymphocytes of normal and leukemia AKR 77-80. mice. Cancer Res. 38, 1113-I 119. CREPPY, E. E., ST~RMER, F. C., R~SCHENTHALER, R., LUSTER, M. I., BCORMAN, G. A.. KORACH, K. S., DIETER, AND DIRHEIMER, G. (1983). Effects of two metabolites M. P., AND HONG, H. L. (1984). Myelotoxicity resulting of ochratoxin A,-(4R)-4-hydroxyochratoxin A and from exogenous estrogens: evidence for bimodal mechochratoxin LY,on immune response in mice. Znfect.Imanism of action. Int. J. Immunopharmacol.. in press. mutt. 39, 1015-1018. DEAN, J. H., PADARATHSINGH, M. L., AND KEYS, L. LUSTER, M. I., DEAN, J. H., AND MOORE, J. A. ( 1982). Evaluation of immune functions in toxicology. In (1978). Response of murine leukemia to combined Methods in Toxicology (W. Hayes, ed.), pp. 56 l-586. BCNV-maleic anhydride-vinyl ether (MVE) adjuvant Raven Press, New York. therapy and correlation with macrophage activation by MVE in the in vitro growth inhibition assay. Cancer MASUDA, E., TAKEMOTO, T., TATSUNO, T., AND OHERA, Treat. Rpt. 62, 1807. T. (1982). Induction of suppressor macrophages in mice DEBRUIN, A. (1976). Interactions with blood corpuscles. by fusarenon-x. Immunology 47, 70 I-708. In Biochemical Toxicity of Environmental Agents (A. PECKHAM, J. C., DOUPNIK, B., AND JONES,0. H. ( I97 1).
Acute toxicity of ochratoxin A and B in chicks. Appl. Microbial.
PITOUT, M. J. (1968). The effect of ochratoxin A on glycogen storage in the rat liver. Toxicol. Appl. Pharmacol. 13,299-306.
PRIOR, M. G., AND SISODIA, C. S. (1982). The effects of ochratoxin A on the immune response of Swiss mice. Canad.
PURCHASE,I. F. H., AND THERON, J. J. (1968). The acute toxicity of ochratoxin A to rats. Food Cosmet. Toxicol. 6,479-483.
PURCHASE, I. F. H., AND VAN DER WATT, J. J. (1971). The long-term toxicity of ochratoxin A to rats. Food Cosmet. Toxicol. 9, 681-682. RICHARD,J. L., THURSTON, J. R., AND PIER, A. C. (1978). Effects of mycotoxin on immunity. In Toxins: Animal, Plant and Microbial (R. Rosenberg, ed.). Pergamon Press, New York. SCOTT, P. M., VAN WALBEEK, W., HARWIG, J., AND FENNELL, D. I. (1970). Occurrence of a mycotoxin, ochratoxin A, in wheat and isolation of ochratoxin A in citrinin producing strains of Penicillium viridicatum. Canad.
J. Plant Sci. 50, 583-585.
SHERIDAN, J. W., METCALF, D., AND STANLEY, E. R. (1975). Further studies on the factor in lung-conditioned medium stimulating granulocyte and monocyte colony formation in vitro. .I Cell. Physiol. 84, 147-158. SHOTWELL, 0. L., HESSELTINE, C. W., AND GOULDER, M. L. ( 1969). Ochratoxin A: Occurrence as natural contaminant of a corn sample. Appl. Microbial. 17, 765766.
SILINI, G., ANDREOZZI, U., AND POZZI, L. V. (1976). The role of the spleen in the repopulation of the haemopoietic system of heavily-irradiated mice. Cell Tissue Kinet. 9, 341-350. SUZUKI, S., KOZUKA, Y., SATOH, T., AND YAMAZAKI, M. (1975). Studies on the nephrotoxicity of ochratoxin A in rats. Toxicol. Appl. Pharmacol. 34, 479-490. SZCZECH,G. M., CARLTON, W. W., AND Turrn, J. (1973a). Ochratoxicosis in Beagle dogs. I. Clinical and clinicopathological features. Vet. Pathol. 10, 135-154. SZCZECH, G. M., CARLTON, W. W., AND TUITE, J. (1973b). Ochratoxicosis in Beagle dogs. II. Pathology. Vet. Pathol. 10, 219-231. TILL, J. E., AND MCCULLOCH, E. A. (1961). A direct measurement on the radiation sensitivity of normal mouse bone marrow cells. Radiat. Res. 14, 213. VAN DER MERWE, K. J., STEYN, P. S., FOURIE,L., SCOTT, DE B., AND THERON, J. J. (1965). Ochratoxin A, a toxic metabolite produced by Aspergillus ochraceus Wilh. Nature (London) 205, 1112-l 113. VAN WALBEEK, W., Scorn, P. M., HARWIG, J., AND LAWRENCE, J. W. (1969). Penicillium viridicatum Westling: A new source of ochratoxin A. Canad. J. Microbial. 15, 1281-1285. VAN WALBEEK, W., MOODIE, C. A., Scorr, P. M., HARWIG, J., AND GRICE, H. C. (197 1). Toxicity and excretion of ochratoxin A in rats intubated with pure ochratoxin A or fed cultures of Penicillium viridicatum. Toxicol. Appl. Pharmacol.