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Metabolic toxicity of simple hydrazines: Monomethyl hydrazine (MMH)
TOXICOLOGY
AND
APPLIED
PHARMACOLOGY
49,
225-236 (I 979)
Metabolic Toxicity of Simple Hydrazines: Monomethyl Hydrazine (MMH)‘12 FRANK N. DOST, D. E. JOHNSON, AND CHIH H. WANG Department
of
Agricultural
Chemistry and Radiation Center, Oregon State University, Corvallis, Oregon 97331
Received August 8, 1978; accepted November
29, 1978
Metabolic Toxicity of Simple Hydrazines: Monomethyl Hydrazine (MMH). Dosr, F. N., JOHNSON, D. E., AND WANG, C. H. (1979). Toxicol. Appl. Pharmacol. 49, 225-236. Radio respirometric data from rats continuously infused with specifically Y-labeled glucose and other substrates indicates that monomethyl hydrazine (MMH) inhibits glycolysis. The effect is evidently at the phosphofructokinase step. MMH was infused SC for I4 hr at 0.036 mmol/kg/hr. Substrates were infused from Hours 4 to 24 through an indwelling intestinal cannula, providing for observation of near steady-state catabolic activity. A dose rate of 0.018 mmol/kg/hr caused a detectable inhibition, and 0.009 mmol/ kg/hr was without effect. The use of continuous substrate infusion techniques and the interpretation of data with respect to metabolic pathway activity is discussed.
The influence of simple hydrazines on carbohydrate metabolism has been acknowledged for more than 60 years, but the mechanisms by which hydrazines alter utilization, synthesis, or storage of carbohydrates are still poorly understood. Smith (1965) investigated the influence of hydrazines on brain slice metabolism in an effort to understand their convulsive effects, and found that monomethyl hydrazine (MMH) increased 14C0, production from [ lJ4C]glucose while decreasing conversion of [2-r4C]- and [614C]glucose. Lactate accumulation and decreased oxygen consumption suggested that pyruvate utilization was inhibited. Amenta 1 Presented in part at the Sixth Annual Conference on Environmental Toxicology, Dayton, Ohio, October 22, 1975. ‘This research was supported principally by contract F 33615-71-C-1516, 6570th Aerospace Medical Research Laboratory, USAF Systems Command, Wright Patterson AFB, Ohio, and in part by Grants ES 00841 and ES 00210, National Institute of Environmental Health Sciences, National Institutes of Health. 225
and Dominguez (1965) observed a decreased oxidation of uniformly labeled [14C]glucose after acute intoxication by relatively high doses of l,l-dimethyl hydrazine (UDMH); MMH was not studied. Hydrazine itself has been studied in somewhat more detail. Hydrazine has been found to increase concentrations of citric acid cycle intermediates, pyruvate, and lactate and to depress levels of triose and hexose phosphates (Fortney et al., 1967; Ray et al., 1970). Ray et al. (1970) suggested that these changes indicated a specific blockade of the acetoacetate-phosphoenolpyruvate transformation, which was supported by finding that phosphonelpyruvate carboxykinase was inhibited in citro by hydrazine. Incorporation of labeled aspartate, pyruvate, and alanine into glucose was also impaired (Fortney et al.., 1967). Interference with gluconeogenesis may be implied from these observations, as well as from the simpler studies of blood glucose and glycogen changes by Izume and Lewis, in 1926. The latter studies were somewhat 0041-008X/79/080225-12%02.00/0 Copyright 0 1979 by Academic Press, Inc. All rights of reproduction in any form reserved. Printed in Great Britain
226
DOST,
JOHNSON,
difficult to interpret because lactate-induced hyperglycemia was amplified by hydrazine, suggesting that a segment of the gluconeogenesis process was either accelerated or compensating for a deficit elsewhere. It is not unreasonable to expect increased glycolytic activity in association with decreased gluconeogenesis, but there is presently no direct evidence that such a shift may be caused by hydrazine, and some work suggests that the opposite effect may occur. Decreased insulin response to glucose loading has been shown following hydrazine treatment (Aleyassine and Lee, 1971; Potter et al., 1969) which may have a secondary effect of slowing glycolysis. Potential effects on citric acid cycle activity have been even less well investigated. Krulik (1966) found that hydrazine interfered with oxidation of cr-keto acids in liver homogenates, but had little effect on succinate oxidation, to some extent paralleling Smith’s (1965) findings after MMH treatment of brain slices. These few specific metabolic investigations, along with the several observations of hydrazine-induced changes in glucose or glycogen dynamics (Underhill, 1911; Underhill and Fine, 1911; Underhill and Hogan, 1915; Izume and Lewis, 1926; Trout, 1966; Fortney, 1966, 1967; Fortney and Clark, 1967; Fortney et al., 1967; Amenta and Johnston, 1962; Taylor, 1966;’ Clark et al., 1970; Potter et al., 1969; Ray et al., 1970) do not permit a satisfactory judgment of the mechanism or significance of toxic effects of simple hydrazines. The sharp divergence between hydrazine and MMH in effects on glucose and glycogen regulation3 suggest in A As examples, in rats at doses of 0.5 LDSO: Hydrazine blocks glycogen synthesis during glucose loading; MMH has no effect. MMH slows mobilization of glycogen from liver when content is more than about 1 y0 liver weight; hydrazine has no effect. In fasted animals, MMH causes hyperglycemia; hydrazine lowers blood glucose. Following a 500-mg glucose load, however, hydrazine sharply decreases glucose tolerance and MMH has no significant effect (unpublished, Dost et al.).
AND
WANG
addition that metabolic effects ascribed to any one member of this class cannot be accepted as representative. Evidence is presented here which indicates that MMH treatment of intact rats causes an inhibition of glycolysis, apparently at the phosphofructokinase step, Activity of the pentose phosphate pathway appeared to be enhanced, presumably secondary to the glycolytic interference; citric acid cycle activity was apparently not affected. The experiments illustrate a radiorespirometric approach, in which various specifically labeled substrates are infused continuously to intact rats through an indwelling intestinal cannula. By infusing a moderate loading of substrate over periods up to 20 hr, a near steady state in catabolism of the substrate was achieved. This procedure partially overcomes compartmental factors which interfere with interpretation of experiments conducted with single administration of labeled material. These stuches are not designed to determine absolute physiological values, although such data can be derived, Rather, the intent is to establish qualitatively whether effects on glucose catabolism occur as a result of toxicity and to learn which specific pathway components are altered. METHODS Male Sprague-Dawley rats obtained from Horton Laboratories, Oakland, California, or from the De: partment of Pharmacology, Oregon State University, were used in all studies reported here. Intestinal cannulas were surgically implanted at animal weights of 28&300 g under Na-pentabarbital anesthesia. At least 5 days of postsurgical rest or return to presurgical weight was required before use. Purina laboratory chow and water ad fibitum were provided at all times other than at surgery and during experimental preparation. Surgical technique. Medical grade vinyl tubing (0.76 x 1.22 mm o.d.) was inserted in the duodenum through a 22-gauge needle perforation about 4-6 mm from the pylorus and passed about 10 mm down the tract. The tube was anchored with 4-O silk to the serosa of the pylorus and passed through the abdominal wall, where it was also anchored, and was then passed forward and upward under the skin to emerge at the back of the neck.
METABOLIC
TOXICITY
OF METHYL
Labeled glucose was administered at a rate of 150 mg and approximately 0.1 .&i/rat/hr through an indwelling duodenal cannula. This input is somewhat less than the carbon turnover by the animal, and very rarely caused blood glucose to exceed 100 mg/ 100 ml. In several experiments, glucose concentration as measured in samples drawn from an indwelling cannula in the posterior vena cava of unanesthetized rats over varying periods of glucose infusions (Table 1). These concentrations are consistent with measurements by Depocas (1964) during similar infusions to rats. Over a period of 20 hr of infusion at this intake rate, liver glycogen remains between 1 and 2% of liver weight, and negligible radioactivity appears in urine. Fructose was infused at a rate of 50 mg/hr accompanied by 100 mg glucose. At higher doses, fructose often caused a lethal toxicity of unknown mechanism in animals receiving MMH. Other substrates were administered at low or trace levels, since their concentrations in blood are low, and were accompanied by unlabeled glucose at 150 mg/rat/hr. The substrate infusion volume was 1.47 ml/hr in all experiments. Other investigators using the concept of continuous infusion of substrates have employed similar or somewhat higher glucose input rates on a body weight basis in the rat (Depocas, 1964), dog (Balasse, 1971), and human (Ceresa et al., 1968; Reaven and Farquhar, 1969). Radioactive substrates. Listed by supplier, position of label, and specific activity in microcuries per micromole: New England Nuclear: glucose, l-, 6.45; glucose, 2-, 2.65; glucose, 3-, 10.0; glucose, 3,4-, 13.89; glucose, 6-, 5.0; Na-pyruvate, l-, 7.35; Napyruvate, 2-, 8.1; Na-acetate, l-, 2.0; Tracerlab: Fructose, 6-, 1.0; Calbiochem: L-Na-aspartate, 1, 2.0; L-Na-glutamate, 1, 3.6.
Radiorespirometry. Respiratory i4C0, from up to four animals was measured individually in l-liter flow ionization chambers coupled to Cary Model 31 vibrating reed electrometers. Output of the electrometers was measured every 3 min by a digital voltmeter coupled to a punched tape recorder (HewlettPackard 2013K data acquisition system). Data were processed by a Hewlett-Packard 9821 programmable calculator with tape reader and plotter. Each experiment was initially calculated every 15 min as the average of five observations during that interval. (The data are shown in Fig. 1.) For general presentation, every fourth such value (hourly) was averaged from two to four experiments. Substrates were diluted to deliver approximately 10 x 103 dpm W/5 min, and specific activity was then accurately determined. All radiorespirometric data was then normalized to the arbitrary standard input: normalized r4C02 output = measured r4C02 output x (10 x 103)/(actual 14C input) all factors expressed as dpm/5 min. This convention enables direct comparison of experiments, and eliminates the need to precisely adjust substrate specific activity to some common value. Animals were held in plastic restraining cages, with the head and shoulders confined in a “helmet” fashioned from a 6-0~ plastic bottle. Room air was drawn forward past the head of each subject at 500 ml/min, through a drying column, ionization chamber, flow meter, metering valve, pump, and exhaust. The detection system was calibrated periodically by continuous injection of a standardized sodium bi [Wlcarbonate solution into 2 N sulfuric acid, from which all emergent “CO, was collected and measured in the same manner as respiratory gases.
TABLE BLWD
GLUCOSE
DURING
227
HYDRAZINE
1
CONTINUOUS
INFUSION
OF
150 mg GLucoss/hr TO 300-g RATS’ No. of samples
Sampling period (min)
Mean concentration (mm/100 ml)
fSD
3 4 3 3 8 6 12
5 8 10 10 25 45 147
80 89.7 69 67.3 98.4 77.2 87.4
2.8 6.8 2.9 5.6 2.9 2.9 7.1
a Sample volume 30 ~1 blood, 10 ,ul plasma was assayed in the Beckman glucose analyzer. Blood glucose calculated on basis of hematocrit, uncorrected for erythrocyte glucose concentration. Animals were unanesthetized.
228
DOST, JOHNSON, AND WANG
All hexoses and amino acids are routinely examined by thin-layer chromatography for purity upon receipt and at any usage 3 months or more following the preceding use. The systems used for hexoses are: (1) t-butanol : methyl ethyl ketone : formic acid : water (8 : 6 : 3 : 3); (2) methanol : pyridine : acetic acid : water (6 : 6 : 1 : 4). The amino acids used here are chromatographed in the first system. Silica Gel G on glass plates is used in each case. Na-pyruvate tends to slowly degrade in storage; this is not significant up to 3 months and the substrate is not used after that period. The oxidation of carbon-4 of glucose is estimated from the difference between the rates of i4COZ production from [3-‘%]glucose and [3,4-i4C]glucose. Administration of MMH. At the beginning of the experimental period, rats which had been fasted 24 hr were placed in restrainers, a 26-gauge needIe without hub was attached to the infusion tubing and emplaced
SC.After an hour of acclimation the infusion was begun. The MMH dose rate for most experiments was 0.036 mmol kg/hr in a volume of 0.067 ml/hr, infused for 14 hr. (Total dose 0.5 mmol/kg, approximately the single administration LDSO for this strain of rats.) In very rare cases this treatment resulted in prodromal or overt signs of acute intoxica-. tion, and data from these animals was not used. The time schedule for each experiment was: time zero, begin MMH infusion; Hour 4, begin “Clabeled substrate infusion; Hour 14, end MMH infusion; Hour 24, end substrate infusion.
RESULTS Untreated Animals
When continually infused into normal animals, each specifically labeled carbon of
FIG. 1. Nature of radiorespirometric data averaged over 15-min intervals during oxidation of [6-Vlglucose by untreated (top) and MMH-treated (bottom) rats. [MMH infused at 0.036 mmol/ kg/hr from time 0 to Hour 14. “‘C-Labeled substrate infused from Hours 4 to 24. Infusion rate normalized to 10 x 103 dpm/5 min (10 on vertical scale).]
METABOLIC
TOXICITY
[14C]glucose was converted to 14COt at a characteristic rate. In every case, there was an initial 4- to 6-hr phase of rapid increase in the 14C0, output followed by a second phase of less rapid increase that often continued till infusion was stopped. Figure 1 shows the general nature of radiorespirometric data from rats, using as
OF METHYL
HYDRAZINE
229
an example the oxidation of [6J4CJglucose by individual treated and untreated animals. The rate of 14C0, output was averaged over 15-min intervals. All other data (Fig. 2) are shown as the mean hourly rates of two or more experiments, with ranges shown periodically. As a reference in examining the radiorespirometric patterns of the various
FIG. 2. Effect of monomethyl hydrazine (MMH) on oxidation to WO, of various labeled metabolic substrates. [Treated animals represented by dashed line. MMH infused at 0.036 mmol/kg/hr from time 0 to Hour 14, W-Labeled substrate infused from Hours 4 to 24. Infusion rate normalized 10 x IOJ dpm/5 min (10 on vertical scale). Curves for carbon 4 of glucose calculated from [3-*4C]glucase and [3,4-%]gIucose data. Curves represent averages of two animals except [3-WJglucose and [6JT]ghcose (control). (3) and [l-W]glucose and [3,4J4C]glucose (intox.) (4). Vertical lines indicate ranges of 14COz production rates.]
230
DOST,
JOHNSON,
continuously infused substrates, the emergence of r4C0, from infused NaH14C0, (Fig. 2, bicarb) is considered to simulate the time base for equilibration of metabolic CO, with endogenous CO, and subsequent release. In normal animals, conversion of [1J4C]-, [2-‘4C]-, and [6J4C]glucose to 14C0, proceeded somewhat similarly, although [6J4C]glucose often was metabolized in a pattern in which the transition between the two phases was not distinct. The rate of oxidation of carbons-3 and -4 increased more rapidly than that of other labels in the early part of infusion and tended to increase a little less rapidly during the second phase of activity. These carbons are much less subject to diversion into carbon sinks which delay ultimate oxidation because they are lost before entry into the citric acid cycle. The generally constant rate of change in i4C0, output through the second phase is presumed to represent movement of glucose carbon into a secondary metabolic pool or pools faster than it is released for oxidation. The slope in each case is sufficiently constant to be represented by a straight line, fitted in this case by inspection. It is reasonable to assume that the zero time intercept of that slope approximates the rate of oxidation above which diversion into a secondary pool occurs. It also therefore shows the extent of direct or “prompt” oxidation of administered glucose carbon, as distinguished from later oxidation of reformed glucose or other products formed from glucose carbon outside the main line of glycolysis and the pentose cycle. The estimated extents of direct oxidation of each glucose carbon are, in order: C-l, 28; C-2, 18; C-3, 36; C-3, 4, 47 ; C-4, 52 (estimated from an indirectly derived slope); C-6, 20% (Fig. 2). A weighted average (accounting for absence of data for C-5) provides an overall fraction of about 31% if the calculated configuration of C-4 activity is used and about 28 % if data from [3, 414C]glucose is used directly instead. These estimates corres-
AND
WANG
pond with the only other estimate recorded under directly comparable conditions, of 30 % (Depocas, 1964). Most estimates of prompt oxidation of glucose in infusion experiments have utilized uniformly labeled glucose, and fall between 15 and 25 “/, depending on the extent of fasting and the rate immediately derived from glucose (Shipley et al., 1970; Balasse, 1971). Substrates which enter the glycolytic sequence later than glucose and which ultimately form labeled acetate oxidize in a pattern similar to that of glucose. [6-i4C]Fructose and Na-[2J4C]pyruvate (Fig. 2), and [I J4C]glycerol (not shown) are examples. Oxidation of the first carbon of pyruvate reaches equilibrium quickly, presumably because access to the decarboxylation reaction is immediate, and proceeds thereafter at a relatively constant rate, as do substrates which enter the citric acid cycle directly, such as Na- [1-14C]glutamate (Fig. 2), and Na[ 1J4C]aspartate and Na-acetate labeled in either position (not shown). The rate of conversion of the respective glucose carbons to 14C0, is depressed to varying degrees in intoxicated animals. It is not possible to assign a regular rate of change in each case which can be extrapolated to time zero, as was done with the control experiments. The time courses have a number of characteristics that may be used in evaluation of the metabolic effect of intoxication, however. In intoxicated animals, output of 14C0, was more or less constant from the first equilibrium at about 6 hr through about 4 hr after the infusion of MMH ended, except in the case of [lJ4C]glucose. Apparently, recovery then began, and the rate of oxidation of all labels except C-6 rose rapidly to approach that of untreated animals by the end of glucose infusion. Even though depressed to some degree, carbons 1, 2, and 3 were converted to 14C02 throughout the experiment at a rate greater than the extrapolated direct oxidation rate ; oxidation of [lJ4C]glucose was decreased only slightly.
METABOLIC
TOXICITY
OF METHYL
DISCUSSION
In contrast, C-4 and C-6 were converted to COZ at a much lower rate than the carbons of the other half of the glucose molecule and turned over at a rate lower than the presumed oxidation rate in normal animals. The effect on [6J4C]fructose (Fig. 2) (and on [lJ4C]frustose, not shown) was of the same order as that of [lJ4C]glucose. Oxidation of Na- [I J4C]acetate and Na- [214C]acetate, Na- [ 1-14C]pyruvate, Na- [214C]pyruvate, Na- [ lJ4C]glutamate, and Na[lJ4C]aspartate were unchanged by MMH. The relationship of the respective apparent rates of conversion of the various glucose labels throughout the time course is shown in Table 2. The rate of 14C0, production from each label by intoxicated animals is tabulated as percentage of the rate in control animals at the same point in the time course. In each case, conversion was faster in treated rats early in the experimental period. Additional experiments with [6-L4C]glucase at lower doses show that 0.018 mmol MMH/kg/hr produced a modest inhibition with a similar time course to that of higher doses, and 0.009 mmol/kg/hr had no effect.
Metabolic pathway derangement in MMHtreated, intact animals is rarely studied, and the use of continuously infused labeled substrates in evaluating metabolic activity is also infrequent. As a consequence, the nature of the system should be reviewed to assist in understanding the toxicologic information. Use of a metabolic map may assist in following the discussion. Experiments in which labeled substrate is administered as a single dose have the limitation that a steady state is never achieved; concentration of substrate or intermediates continuously changes in all compartments. This condition may mask changes in rate or extent of reactions which have been induced by toxicity. The value of continuous infusion of substrates was established long ago (Coxon and Robinson, 1959); Shipley et al. (1970).recognized that the approach could dispense with the complex models needed for single dose experiments. Shipley et al. (1970) also suggested, however, that long experiments might compromise the subject and this concern has limited such work to periods of 2-5 hr, usually dealing with regulatory mecha-
TABLE EFFECT
OF MONOMETHYL
HYDRAZINE
231
HYDRAZINE
(MMH)
2 ON
14C0,
FORMATION
FROM
VARIOUS
LABELS
OF [“+C]GLUCOSE
Hours after beginning MMH Begin MMH 4 0
2
Glucose [I-T] [2-‘4CJ [3-Y 1 [3,4-W] 4b [6-W]
Begin glucose 4 4 6
End MMH J 14
16
18
20
22
19 75 108 76 50 40
87 86 107 86
98 93 123 91
45
60
8
10
12
94 70
81 55
75 55
91
88
74
78
loo
86
41 125
75 80
95 110 86 76 69 79
97 100
138
105 130 102
74 55 62
81 74 46
58 47 34
57 30 30
145”
End glucose 4 24
a WO, output, expressed as percentage of output by untreated rats. ’ Calculated as ([(3,4-Y-(3-W)] intox}/{[(3,4-L4C)-(3-‘4C)] control}, where each factor is the respiratory WO, as net counts per minute per 5 min arising from the respective labeled glucose at time (1).
232
DOST,
JOHNSON,
nisms rather than metabolism. We have applied the concept to make a semiquantitative evaluation of the impact of intoxicants on the main avenues of carbohydrate metabolism in intact animals. It is clear from the data that at least 4 to 6 hr of observation is necessary before the impact of intoxication on rates of metabolic activity can be assessed, and 24 hr of observation are needed for full evaluation. The few other investigators who have used continuous infusion of more than traces of glucose have also found a similar delay, but the dose rates and general methods differ so extensively that comparison is difficult (Stretten et al., 1951; Wick et al., 1951; de Freitas and Depocas, 1965; Balasse, 1971; Depocas, 1959; Searle et al., 1956; Bergman, 1963). The biphasic nature of the rate plots for 14C0, output suggests at least two major influences on the time required to achieve a steady state. The most probable governing factors are the rates of release of label from various metabolic pools or sinks. Physical dilution of infused iabeled glucose in the glucose compartments of the body is apparently very fast (Shipley et al., 1967; Searle et al., 1956) and does not contribute to the delay. A comparison by Baker et al. (1959) of extracellular. glucose space compared with total tissue glucose as measured by other investigators (Cori et al., 1953; Orten and Sayers, 1942; Gey, 1956), shows that virtually no glucose exists as such within cells, which means that glucose is diluted only in the extracellular space. Equilibration of metabolic CO, also probably has little influence on the time course. When bi[14C]carbonate is infused continuously, 14C0, output becomes constant as a rate between 90 and 95 % of input, within about an hour (Fig. 2), which is probably equivalent to the distribution and mixing time of metabolic CO,. A similar time course with a more pronounced second phase has been found by Morris and Simpson-Morgan (1963) over a 3-hr period. In theory, randomization of infused glucose to glucose labeled in other positions
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WANG
could contribute to the extended period required to reach a steady state, but such effect should be minimal in our studies. Pentose cycle activity is minimal and does not contribute to unequal flow of C-l and C-6 into acetate-derived products. The glucose loading of these experiments should limit gluconeogenic flow of “relabeled” glucose. Available evidence indicates that loading does inhibit reflux of glucose (Parli et al., 1969). In dogs (Cowan et al., 1969) and rabbit liver (Williams et al., 1971), loading of glucose has been shown to maintain C-l/C-6 ratios in 14C02 at about 1, and to prevent endogenous glucose production (Long et al., 1971; Hetenyi and Wrenshall, 1968; von Holt et al., 1961). There are considerable data to indicate that there are extensive metabolic sinks or pools into which carbon enters, no longer as a constituent of glucose. A single dose of labeled glucose disappears from plasma within minutes, transformed into glucose intermediates; labeled CO, does not appear in significant amounts during this early phase (Baker et al., 1959; Baker and Huebotter, 1964). Shipley et al. (1970) have discussed the great capacity of metabolic pools, particularly lipid compartments for glucose carbon prior to oxidation. Whether considered as slowly releasing pools such as fat, or accumulation of more rapidly processed metabolites in the direct glycolytic sequence, when removal is slower than intake the rate of removal becomes rate limiting for release of 14C0 As substrate (or”’ intermediate) builds up there should be a gradual constant increase in the rate of output of 14C0, until some new limiting step is reached or until the entire system is at equilibrium with intake. It seems reasonable to attribute the second phase of gradual and constant increase in rate of 14C0, output to such a series of reactions. The first phase of more rapid change must reflect a period of approach to constant rate among the series of more immediately accessible reactions.
METABOLIC
TOXICITY
Evidence in our experiments supports the contention that the equilibration periods are functions of the time required for carbons to complete reaction sequences. [6-r4C]Fructose and Na- [2J4C]pyruvate which label acetate, require the same period of equilibration as does glucose (Fig. 2), but Na-[1-*4C]pyruvate, which is decarboxylated before any diversion to nonglucose products can occur, is converted very rapidly, and has onIy a limited slope in the second phase. Since Na[ 1J4C]glutamate and Na- [ 1-14C]aspartate are known to enter the citric acid cycle readily and they are immediately oxidized, it may be implied that the cycle moves quickly and is not linked with a metabolic sink for carbon. In view of the respective behavior of acetatelabeling and pyruvate-l-labeling precursors it is fair to assume that flux of acetate into fatty acid synthesis and oxidation temporarily disposes of a large amount of glucose carbon. Oxidation of glucose, fructose, and [2J4C]pyruvate are all similarly delayed, but oxidation of [I-r4C]pyruvate is rapid. The difference between the relatively slow turnover of C-3 of glucose, which eventually becomes carbon-l of pyruvate, and the rapid oxidation of exogenous [lJ4C]pyruvate can be explained only by the time required for biochemical equilibration of glucose at other points in the sequence. The times required for overall isotopic equilibration as judged by 14C0, production are more or less constant at infusion rates from traces of [r4C]glucose up to 500 mg/hr. Furthermore, if we infuse 150 mg unlabeled glucose/hr to rats under the conditions of experiments reported here and after 6 hr add a trace label to the continued glucose infusion, the expected 4- to 6-hr elapse before the first equilibration phase as represented by 14C0, output is completed. A single priming dose of labeled material superimposed on the system in an effort to establish an earlier steady state will influence all compartments unequally for about the same period and provides no advantage in continuous infusion experiments.
OF METHYL
HYDRAZINE
233
Changes in 14C02 output resulting from acute intoxication during infusion are not subject to the same delays because all reactions have reached maximum rates, dependent either on availability of substrate or reaction capacity. Animals with previously established toxicity are subject to the same kinds of time course, although specific rates within the system may be changed, and the most useful information is the rate data obtained after the first phase is complete. Acute non lethal fluoride intoxication, for example, which apparently blocks glucose entry to cells while causing no obstruction in main pathways, results in an almost immediate drop in 14C0, production from all [14C]glucose labels (Dost et al., 1977). Interpretation of Changes Resulting from Intoxication In the process of interpreting observed toxic effects it is first necessary to establish whether the change is in the direct oxidation of glucose (glycolysis, pentose cycle, and the citric acid cycle) or on biosynthetic pathways into which carbon may move for later return and oxidation. If MMH primarily influenced movement of carbon into or out of reaction compartments not in the direct line of metabolism, a much greater symmetry of effect on the two halves of the glucose molecule should be evident. We should also see an appreciable difference between oxidation rates for C-3 when compared with C-2, and for C-4 compared with C-6. The sharp singular decrease in C-6 oxidation and the decrease of C-4 and C-6 oxidation below the level of estimated direct metabolism indicates oxidation of the C-4, 5, 6 half of the molecuIe is selectively obstructed, which can only happen in a main pathway, probably glycolysis. The rate of conversion of continuously infused [6J4C]glucose to 14C0, will represent, within limits, the total glycolytic flux because pentose cycle activity is normally slight and the glucuronic acid pathway is negligible. A small additional fraction of the 14C0, from
234
DOST,
JOHNSON,
C-6 will arise from glucose cycled through the pentose pathway; of every three glucose molecules entering that pathway, one terminal 3-carbon fragment (C-4, 5, 6) will enter triose metabolism as glyceraldehyde-3-phosphate. MMH caused a marked decrease in production of 14C0, from [6J4C]glucose and a similar apparent decrease in conversion of carbon-4. These changes indicate glycolytic impairment, since the 4,5,6-carbon fragment must remain intact through either glycolysis or the pentose cycle until carbon 4 is lost in decarboxylation of pyruvate. The rate of conversion of [6-r4C]glucose under those conditions will then represent the amount of intact glucose passing through glycolysis, including that reformed during pentose cycle activity, and less that arising from glyceraldehyde-3-phosphate which was formed from a terminal 3-carbon unit in the pentose pathway. Differentiation of these two components is difficult if not impossible to precisely measure in an intact animal system. Carbons 1, 2, and 3 of glucose may be assumed to traverse glycolysis proper to the same extent as carbon 6. The extent of C-l oxidation in excess of C-6 oxidation may be taken as an indication of increased activity of the pentose pathway. A similar effect of acetyl phenylhydrazine on glycolysis has been observed in erythrocytes (Szeinberg and Marks, 1961; Kosower et al., 1963; Mills and Buell, 1965). Having established that glycolysis was impaired, it was necessary to learn where in this rather extensive sequence the defect might be. (The possibility of interference in the citric acid cycle was considered unlikely since oxidation of Na- [ 1J4C]pyruvate and Na- [2J4C]pyruvate, and Na [ lJ4C]aspartate, and Na- [ 1J4C]glutamate were all unaffected by MMH.) The next step was to determine whether the effect was on hexose or triose metabolism. Fructose is the best substrate for this problem because it enters glycolysis rapidly in normal
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WANG
animals as glyceraldehyde phosphate and dihydroxyacetone phosphate. The limited decrease in [6-14C]fructose catabolism under MMH intoxication indicates that consumption of triose was not seriously slowed, and directed attention to hexose catabolism. (Use of [lJ4C]fructose provided identical information but is not reported.) On the basis of the considerably greater conversion to 14C0, of labeled C-2 and C-3 than C-6, some aspect of hexose metabolism is clearly capable of proceeding. This can only occur if fructose 6-phosphate formed in the pentose cycle was readily transformed to glucose 6-phosphate in the bidirectional phosphoglucose isomerase reaction. Since the fructaldolase conversion of fructose 1,6-phosphate to trioses was not impaired it is reasonable to assume that fructose 1,6phosphate metabolism to trioses was also unaffected since the same enzyme is required. The remaining step and therefore the logical site where inhibition must occur is the key regulatory phosphofructokinase reaction.
REFERENCES H., AND LEE, S. H. (1971). Inhibition by hydrazine, phenelzine and pargyline of insulin release from rat pancreas. Endocrinology 89, 125129. AMENTA, J. S., AND JOHNSTON, E. H. (1962). Hydrazineinduced alterations in rat liver. Lab. Invest. 11, ALEYASSINE,
956-962. AMENTA, J. S., AND DOMINGUEZ,
A. M. (1965). The effect of hydrazine and congeners on Ci403 respiratory pattern of various metabolic substrates. Toxicol. Appl. Pharmacol. I, 236-246. BAKER, N., SHIPLEY, R. A., CLARK, R. E., AND INCEFY, G. E. (1959). Cl4 studies in carbohydrate metabolism: glucose pool size and rate of turnover in the normal rat. Amer. J. Physiol. 196, 245-252. BAKER, N., AND HUEBO~ER, R. (1964). Glucose metabolism in mice. Amer. J. Physiol. 207, 11551160. BALASSE, E. 0. (1971). Effect of free fatty acids and ketone bodies on glucose uptake and oxidation in the dog. Norm. Metab. Res. 3, 403-409. BERGMAN, E. N. (1963). Quantitative aspects of glucose metabolism in pregnant and nonpregnant sheep. Amer. J. Physiol. 204, 147-152.
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F., GHEMI, F., MARTINI, P. F., MARTINO, P., SEGRE, G., AND VITELLI, A. (1968). Control of blood glucose in normal and diabetic subjects. Studies by compartment analysis and digital computer techniques. Diabetes 17, 570-578. CLARK, D. A., LEEDER, L. G., FOULDS, E. L., AND TROUT, D. L. (1970). Changes in lipids of rat liver after hydrazine injection. &o&em. Pharmacol. 19, 1743-1752. CORI, G., Cross, J., AND CORI, C. (1953). Fermentable sugar in heart and skeletal muscle. J. Biol. Chem. CERESA,
103, 13-24.
COWAN, J. S., SCHACHTER,D., AND HETENYI, G., JR. (1969). Validity of a tracer-injection method for studying glucose turnover in normal dogs. J. Nucl.
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10, 98-102.
COXON, R. V., AND ROBINSON, R. J. (1959). Movements of radioactive carbon dioxide within the animal body during oxidation of Y-labeled substances. J. Physiol. 147, 487-510. DEFREITAS, A. S. W., AND DEPOCAS, F. (1965). Fatty acid and glyceride glycerol synthesis from glucose during high rates of glucose uptake in the intact rat. Cunud.
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43,437450.
DEPOCAS, F. (1959). The pool size turnover rate, and oxidation rate of body glucose in anesthetized warm- and cold-acclimated rats exposed to a warm environment. Cunad. J. Biochem. Physiol. 37,285-295. DEPOCAS, F. (1964). Regulation of glucose oxidation in the white rat. Amer. J. Physiol. 206, 113-118. DOST, F. N., KNAUS, R., JOHNSON,D., AND WANG, C. (1977). Fluoride impairment of glucose utilizationnature of effect in rats during and after continuous NaF infusion. Toxicol. Appl.Phurmacol.41,451-458. FORTNEY, S. R. (1966). Effect of hydrazine on liver glycogen, arterial glucose, lactate, pyruvate and acid-base balance in the anesthetized dog. J. Pharmacol.
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153, 562-568.
FORTNEY, S. R. (1967). Effect of hydrazine on carbohydrate metabolism in vivo and in vitro. Aerospace Med. 38, 727-731. FORTNEY, S. R., CLARK, D. A., AND STEIN, E. (1967). Inhibition of gluconeogenesis by hydrazine administration in rats. J. Pharmucol. Exp. Ther. 156, 277284. FORTNEY, S. R., AND CLARK, D. A. (1967). Effect of monomethyl hydrazine on methemoglobin production in vitro and in vivo. Aerospace Med. 38, 239-242. GEY, K. (1956). The concentration of glucose in rat tissues. Biochem. J. 64, 145-150. HETENYI, G., JR., AND WRENSHALL, G. A. (1968). Adaptive changes in rates of appearance and disappearance of glucose in dogs following step changes in the rate of glucose infusion. Canud. J. Physiol. Phurmacol. 46, 391-398.
OF METHYL
235
HYDRAZINE
IZUME, S., AND LEWIS, H. B. (1926). The influence of hydrazine and its derivatives on metabolism. J. Biol.
Chem.
71, 51-66.
KOSOWER, N. S., VANDERHOGG, A., JAFFB, R., AND LONDON, M. (1963). Metabolic changes in normal and glucose-6-phosphate dehydrogenase-deficient erythrocytes induced by acetylphenylhydrazine. J. C/in. Invest. 42, 1025-1030. KRULIK, R. (1966). Der Einfluss von hydrazin und Isonicotinslurehydrazid auf bestimmte Substanzen des zuckerstoffwechsels in vitro. Arz. Frosch. 16, 1623-1626. LONG, C. L., SPENCER, J. L., KINNEY, J. M., AND GEIGER, J. W. (1971). Carbohydrate metabolism in normal man and effect of glucose infusion. J. Appl. Physiol.
31, 102-109.
MILLS, G. C., AND BUELL, W. F. (1965). Effects of acetylphenylhydrazine on erythrocyte metabolism. Clin.
Chim.
Actu
11, 310-320.
MORRIS, B., AND SIMPSON-MORGAN, M. W. (1963). The excretion of YO, during the continuous intravenous infusion of NaH%O, in unanesthetized rats. J. Physiol. 169, 713-728. ORTEN, J., AND SAYERS,G. (1942). A comparison of the disposition of injected glucose in two strains of rats. J. Biol. Chem. 145, 123-127. PARLI, C. J., BURNS, L. C., MEGHAL, S. K., KINNEBERG, K., O’NEAL, R. M., AND KOEPPE, R. E. (1969). Pyruvate metabolism in mammalian liver. Amer. POTTER,
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216, 1525-1533.
W. Z., ZAHARKO, D. S., AND BECK, L. V. (1969). Possible role of hydrazine group in hypoglycemia associated with the use of certain monoamine-oxidase inhibitors (MAOI’s). Diabetes 18, 538-541.
RAY, P. D., HANSON, R. L., AND LARDY, H. A. (1970). Inhibition by hydrazine of gluconeogenesis in the rat. J. Biol. Chem. 245, 690-696. REAVEN, G. M., AND FARQUHAR, J. W. (1969). Steady state plasma insulin response to continuous glucose infusion in normal and diabetic subjects. Diabetes 18, 273-279.
SEARLE, G. L., STRISOWER, E. H., AND CHAIKOFF, I. L. (1956). Determination of rates of glucose oxidation in normal and diabetic dogs by a technique involving continuous injection of C14glucose. Amer. J. Physiol. 185, 589-594. SHIPLEY, R. A., CHUDZIK, E. B., GIBBONS, A. P., JONGEDYK, K., AND BRUMMOND, D. 0. (1967). Rate of glucose transformation in the rat by wholebody analysis after g1ucose-‘4C. Amer. J. Physiol. 213, 1149-l
158.
SHIPLEY, R. A., CHUDZIK, E. B., AND GIBBONS, A. P. (1970). Rate of disposal of glucose carbon to CO,, fat, protein, and glycogen in the diabetic rat. Amer. J. Physiol.
219, 364-373,
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SMITH, T. E. (1965). Brain carbohydrate metabolism during hydrazine toxicity. Biochem. Pharmacol. 14, 979-988. STRETTEN, D., JR., WELT, I. D., INGLE, D. J., AND MORLEY, E. H. (1951). Rates of glucose production and oxidation in normal and diabetic rats. J. Biol. Chem. 192, 817-830. SZEINBERG, A., AND MARKS, P. A. (1961). Substances stimulating glucose catabolism by the oxidative reactions of the pentose phosphate pathway in human erythrocytes. J. C/in. Invest. 40, 914-924. TAYLOR, G. D. (1966). Efects of Hydra&e on Blood Glucose and Muscle and Liver Glycogen in the Anesthetized Dog. USAF School of Aerospace Med. SAM-TR 66-12. TROUT, D. L. (1966). Effects of hydrazine on fat transport as affected by blood glucose concentration. J. Pharmacol. Exp. Ther. 152, 529-534. UNDERHILL, F. P. (191 I). Studies in carbohydrate metabolism: IV. Do hydrazine derivatives show the
AND
WANG
typical hydrazine effect upon J. Biol. Chem. 10, 159-168.
blood
sugar
content?
UNDERHILL, F. P., AND FINE, M. S. (I91 I). Studies in carbohydrate metabolism. II. The prevention and inhibition of pancreatic diabetes. J. Biol. Chem. 10, 271-283. UNDERHILL, F. P., AND HOGAN, A. G. (1915). Studies in carbohydrate metabolism. VII!. The influence of hydrazine on the utilization of dextrose. J. Biol. Chem. 20, 203-2 15. VON HOLT, C., SCHMIDT, H., FELDMANN, H., AND HALLMANN, I. (1961). Der umsatz der Blutglucose. Biochem. Zeitschrift 334, 524-533, WICK, A. N., DRURY, D. R., BANCROFT, R. W., AND MACKAY, E. M. (1951). Action of insulin on the extrahepatic tissues. J. Biol. Chem. 188, 241-249. WILLIAMS, J. F., RIENITS, K. G., SCHOFIELD, P. J., AND CLARK, M. G. (1971). The pentose phosphate pathway in rabbit liver. Biochem. J. 123, 923-943.
AND
APPLIED
PHARMACOLOGY
49,
225-236 (I 979)
Metabolic Toxicity of Simple Hydrazines: Monomethyl Hydrazine (MMH)‘12 FRANK N. DOST, D. E. JOHNSON, AND CHIH H. WANG Department
of
Agricultural
Chemistry and Radiation Center, Oregon State University, Corvallis, Oregon 97331
Received August 8, 1978; accepted November
29, 1978
Metabolic Toxicity of Simple Hydrazines: Monomethyl Hydrazine (MMH). Dosr, F. N., JOHNSON, D. E., AND WANG, C. H. (1979). Toxicol. Appl. Pharmacol. 49, 225-236. Radio respirometric data from rats continuously infused with specifically Y-labeled glucose and other substrates indicates that monomethyl hydrazine (MMH) inhibits glycolysis. The effect is evidently at the phosphofructokinase step. MMH was infused SC for I4 hr at 0.036 mmol/kg/hr. Substrates were infused from Hours 4 to 24 through an indwelling intestinal cannula, providing for observation of near steady-state catabolic activity. A dose rate of 0.018 mmol/kg/hr caused a detectable inhibition, and 0.009 mmol/ kg/hr was without effect. The use of continuous substrate infusion techniques and the interpretation of data with respect to metabolic pathway activity is discussed.
The influence of simple hydrazines on carbohydrate metabolism has been acknowledged for more than 60 years, but the mechanisms by which hydrazines alter utilization, synthesis, or storage of carbohydrates are still poorly understood. Smith (1965) investigated the influence of hydrazines on brain slice metabolism in an effort to understand their convulsive effects, and found that monomethyl hydrazine (MMH) increased 14C0, production from [ lJ4C]glucose while decreasing conversion of [2-r4C]- and [614C]glucose. Lactate accumulation and decreased oxygen consumption suggested that pyruvate utilization was inhibited. Amenta 1 Presented in part at the Sixth Annual Conference on Environmental Toxicology, Dayton, Ohio, October 22, 1975. ‘This research was supported principally by contract F 33615-71-C-1516, 6570th Aerospace Medical Research Laboratory, USAF Systems Command, Wright Patterson AFB, Ohio, and in part by Grants ES 00841 and ES 00210, National Institute of Environmental Health Sciences, National Institutes of Health. 225
and Dominguez (1965) observed a decreased oxidation of uniformly labeled [14C]glucose after acute intoxication by relatively high doses of l,l-dimethyl hydrazine (UDMH); MMH was not studied. Hydrazine itself has been studied in somewhat more detail. Hydrazine has been found to increase concentrations of citric acid cycle intermediates, pyruvate, and lactate and to depress levels of triose and hexose phosphates (Fortney et al., 1967; Ray et al., 1970). Ray et al. (1970) suggested that these changes indicated a specific blockade of the acetoacetate-phosphoenolpyruvate transformation, which was supported by finding that phosphonelpyruvate carboxykinase was inhibited in citro by hydrazine. Incorporation of labeled aspartate, pyruvate, and alanine into glucose was also impaired (Fortney et al.., 1967). Interference with gluconeogenesis may be implied from these observations, as well as from the simpler studies of blood glucose and glycogen changes by Izume and Lewis, in 1926. The latter studies were somewhat 0041-008X/79/080225-12%02.00/0 Copyright 0 1979 by Academic Press, Inc. All rights of reproduction in any form reserved. Printed in Great Britain
226
DOST,
JOHNSON,
difficult to interpret because lactate-induced hyperglycemia was amplified by hydrazine, suggesting that a segment of the gluconeogenesis process was either accelerated or compensating for a deficit elsewhere. It is not unreasonable to expect increased glycolytic activity in association with decreased gluconeogenesis, but there is presently no direct evidence that such a shift may be caused by hydrazine, and some work suggests that the opposite effect may occur. Decreased insulin response to glucose loading has been shown following hydrazine treatment (Aleyassine and Lee, 1971; Potter et al., 1969) which may have a secondary effect of slowing glycolysis. Potential effects on citric acid cycle activity have been even less well investigated. Krulik (1966) found that hydrazine interfered with oxidation of cr-keto acids in liver homogenates, but had little effect on succinate oxidation, to some extent paralleling Smith’s (1965) findings after MMH treatment of brain slices. These few specific metabolic investigations, along with the several observations of hydrazine-induced changes in glucose or glycogen dynamics (Underhill, 1911; Underhill and Fine, 1911; Underhill and Hogan, 1915; Izume and Lewis, 1926; Trout, 1966; Fortney, 1966, 1967; Fortney and Clark, 1967; Fortney et al., 1967; Amenta and Johnston, 1962; Taylor, 1966;’ Clark et al., 1970; Potter et al., 1969; Ray et al., 1970) do not permit a satisfactory judgment of the mechanism or significance of toxic effects of simple hydrazines. The sharp divergence between hydrazine and MMH in effects on glucose and glycogen regulation3 suggest in A As examples, in rats at doses of 0.5 LDSO: Hydrazine blocks glycogen synthesis during glucose loading; MMH has no effect. MMH slows mobilization of glycogen from liver when content is more than about 1 y0 liver weight; hydrazine has no effect. In fasted animals, MMH causes hyperglycemia; hydrazine lowers blood glucose. Following a 500-mg glucose load, however, hydrazine sharply decreases glucose tolerance and MMH has no significant effect (unpublished, Dost et al.).
AND
WANG
addition that metabolic effects ascribed to any one member of this class cannot be accepted as representative. Evidence is presented here which indicates that MMH treatment of intact rats causes an inhibition of glycolysis, apparently at the phosphofructokinase step, Activity of the pentose phosphate pathway appeared to be enhanced, presumably secondary to the glycolytic interference; citric acid cycle activity was apparently not affected. The experiments illustrate a radiorespirometric approach, in which various specifically labeled substrates are infused continuously to intact rats through an indwelling intestinal cannula. By infusing a moderate loading of substrate over periods up to 20 hr, a near steady state in catabolism of the substrate was achieved. This procedure partially overcomes compartmental factors which interfere with interpretation of experiments conducted with single administration of labeled material. These stuches are not designed to determine absolute physiological values, although such data can be derived, Rather, the intent is to establish qualitatively whether effects on glucose catabolism occur as a result of toxicity and to learn which specific pathway components are altered. METHODS Male Sprague-Dawley rats obtained from Horton Laboratories, Oakland, California, or from the De: partment of Pharmacology, Oregon State University, were used in all studies reported here. Intestinal cannulas were surgically implanted at animal weights of 28&300 g under Na-pentabarbital anesthesia. At least 5 days of postsurgical rest or return to presurgical weight was required before use. Purina laboratory chow and water ad fibitum were provided at all times other than at surgery and during experimental preparation. Surgical technique. Medical grade vinyl tubing (0.76 x 1.22 mm o.d.) was inserted in the duodenum through a 22-gauge needle perforation about 4-6 mm from the pylorus and passed about 10 mm down the tract. The tube was anchored with 4-O silk to the serosa of the pylorus and passed through the abdominal wall, where it was also anchored, and was then passed forward and upward under the skin to emerge at the back of the neck.
METABOLIC
TOXICITY
OF METHYL
Labeled glucose was administered at a rate of 150 mg and approximately 0.1 .&i/rat/hr through an indwelling duodenal cannula. This input is somewhat less than the carbon turnover by the animal, and very rarely caused blood glucose to exceed 100 mg/ 100 ml. In several experiments, glucose concentration as measured in samples drawn from an indwelling cannula in the posterior vena cava of unanesthetized rats over varying periods of glucose infusions (Table 1). These concentrations are consistent with measurements by Depocas (1964) during similar infusions to rats. Over a period of 20 hr of infusion at this intake rate, liver glycogen remains between 1 and 2% of liver weight, and negligible radioactivity appears in urine. Fructose was infused at a rate of 50 mg/hr accompanied by 100 mg glucose. At higher doses, fructose often caused a lethal toxicity of unknown mechanism in animals receiving MMH. Other substrates were administered at low or trace levels, since their concentrations in blood are low, and were accompanied by unlabeled glucose at 150 mg/rat/hr. The substrate infusion volume was 1.47 ml/hr in all experiments. Other investigators using the concept of continuous infusion of substrates have employed similar or somewhat higher glucose input rates on a body weight basis in the rat (Depocas, 1964), dog (Balasse, 1971), and human (Ceresa et al., 1968; Reaven and Farquhar, 1969). Radioactive substrates. Listed by supplier, position of label, and specific activity in microcuries per micromole: New England Nuclear: glucose, l-, 6.45; glucose, 2-, 2.65; glucose, 3-, 10.0; glucose, 3,4-, 13.89; glucose, 6-, 5.0; Na-pyruvate, l-, 7.35; Napyruvate, 2-, 8.1; Na-acetate, l-, 2.0; Tracerlab: Fructose, 6-, 1.0; Calbiochem: L-Na-aspartate, 1, 2.0; L-Na-glutamate, 1, 3.6.
Radiorespirometry. Respiratory i4C0, from up to four animals was measured individually in l-liter flow ionization chambers coupled to Cary Model 31 vibrating reed electrometers. Output of the electrometers was measured every 3 min by a digital voltmeter coupled to a punched tape recorder (HewlettPackard 2013K data acquisition system). Data were processed by a Hewlett-Packard 9821 programmable calculator with tape reader and plotter. Each experiment was initially calculated every 15 min as the average of five observations during that interval. (The data are shown in Fig. 1.) For general presentation, every fourth such value (hourly) was averaged from two to four experiments. Substrates were diluted to deliver approximately 10 x 103 dpm W/5 min, and specific activity was then accurately determined. All radiorespirometric data was then normalized to the arbitrary standard input: normalized r4C02 output = measured r4C02 output x (10 x 103)/(actual 14C input) all factors expressed as dpm/5 min. This convention enables direct comparison of experiments, and eliminates the need to precisely adjust substrate specific activity to some common value. Animals were held in plastic restraining cages, with the head and shoulders confined in a “helmet” fashioned from a 6-0~ plastic bottle. Room air was drawn forward past the head of each subject at 500 ml/min, through a drying column, ionization chamber, flow meter, metering valve, pump, and exhaust. The detection system was calibrated periodically by continuous injection of a standardized sodium bi [Wlcarbonate solution into 2 N sulfuric acid, from which all emergent “CO, was collected and measured in the same manner as respiratory gases.
TABLE BLWD
GLUCOSE
DURING
227
HYDRAZINE
1
CONTINUOUS
INFUSION
OF
150 mg GLucoss/hr TO 300-g RATS’ No. of samples
Sampling period (min)
Mean concentration (mm/100 ml)
fSD
3 4 3 3 8 6 12
5 8 10 10 25 45 147
80 89.7 69 67.3 98.4 77.2 87.4
2.8 6.8 2.9 5.6 2.9 2.9 7.1
a Sample volume 30 ~1 blood, 10 ,ul plasma was assayed in the Beckman glucose analyzer. Blood glucose calculated on basis of hematocrit, uncorrected for erythrocyte glucose concentration. Animals were unanesthetized.
228
DOST, JOHNSON, AND WANG
All hexoses and amino acids are routinely examined by thin-layer chromatography for purity upon receipt and at any usage 3 months or more following the preceding use. The systems used for hexoses are: (1) t-butanol : methyl ethyl ketone : formic acid : water (8 : 6 : 3 : 3); (2) methanol : pyridine : acetic acid : water (6 : 6 : 1 : 4). The amino acids used here are chromatographed in the first system. Silica Gel G on glass plates is used in each case. Na-pyruvate tends to slowly degrade in storage; this is not significant up to 3 months and the substrate is not used after that period. The oxidation of carbon-4 of glucose is estimated from the difference between the rates of i4COZ production from [3-‘%]glucose and [3,4-i4C]glucose. Administration of MMH. At the beginning of the experimental period, rats which had been fasted 24 hr were placed in restrainers, a 26-gauge needIe without hub was attached to the infusion tubing and emplaced
SC.After an hour of acclimation the infusion was begun. The MMH dose rate for most experiments was 0.036 mmol kg/hr in a volume of 0.067 ml/hr, infused for 14 hr. (Total dose 0.5 mmol/kg, approximately the single administration LDSO for this strain of rats.) In very rare cases this treatment resulted in prodromal or overt signs of acute intoxica-. tion, and data from these animals was not used. The time schedule for each experiment was: time zero, begin MMH infusion; Hour 4, begin “Clabeled substrate infusion; Hour 14, end MMH infusion; Hour 24, end substrate infusion.
RESULTS Untreated Animals
When continually infused into normal animals, each specifically labeled carbon of
FIG. 1. Nature of radiorespirometric data averaged over 15-min intervals during oxidation of [6-Vlglucose by untreated (top) and MMH-treated (bottom) rats. [MMH infused at 0.036 mmol/ kg/hr from time 0 to Hour 14. “‘C-Labeled substrate infused from Hours 4 to 24. Infusion rate normalized to 10 x 103 dpm/5 min (10 on vertical scale).]
METABOLIC
TOXICITY
[14C]glucose was converted to 14COt at a characteristic rate. In every case, there was an initial 4- to 6-hr phase of rapid increase in the 14C0, output followed by a second phase of less rapid increase that often continued till infusion was stopped. Figure 1 shows the general nature of radiorespirometric data from rats, using as
OF METHYL
HYDRAZINE
229
an example the oxidation of [6J4CJglucose by individual treated and untreated animals. The rate of 14C0, output was averaged over 15-min intervals. All other data (Fig. 2) are shown as the mean hourly rates of two or more experiments, with ranges shown periodically. As a reference in examining the radiorespirometric patterns of the various
FIG. 2. Effect of monomethyl hydrazine (MMH) on oxidation to WO, of various labeled metabolic substrates. [Treated animals represented by dashed line. MMH infused at 0.036 mmol/kg/hr from time 0 to Hour 14, W-Labeled substrate infused from Hours 4 to 24. Infusion rate normalized 10 x IOJ dpm/5 min (10 on vertical scale). Curves for carbon 4 of glucose calculated from [3-*4C]glucase and [3,4-%]gIucose data. Curves represent averages of two animals except [3-WJglucose and [6JT]ghcose (control). (3) and [l-W]glucose and [3,4J4C]glucose (intox.) (4). Vertical lines indicate ranges of 14COz production rates.]
230
DOST,
JOHNSON,
continuously infused substrates, the emergence of r4C0, from infused NaH14C0, (Fig. 2, bicarb) is considered to simulate the time base for equilibration of metabolic CO, with endogenous CO, and subsequent release. In normal animals, conversion of [1J4C]-, [2-‘4C]-, and [6J4C]glucose to 14C0, proceeded somewhat similarly, although [6J4C]glucose often was metabolized in a pattern in which the transition between the two phases was not distinct. The rate of oxidation of carbons-3 and -4 increased more rapidly than that of other labels in the early part of infusion and tended to increase a little less rapidly during the second phase of activity. These carbons are much less subject to diversion into carbon sinks which delay ultimate oxidation because they are lost before entry into the citric acid cycle. The generally constant rate of change in i4C0, output through the second phase is presumed to represent movement of glucose carbon into a secondary metabolic pool or pools faster than it is released for oxidation. The slope in each case is sufficiently constant to be represented by a straight line, fitted in this case by inspection. It is reasonable to assume that the zero time intercept of that slope approximates the rate of oxidation above which diversion into a secondary pool occurs. It also therefore shows the extent of direct or “prompt” oxidation of administered glucose carbon, as distinguished from later oxidation of reformed glucose or other products formed from glucose carbon outside the main line of glycolysis and the pentose cycle. The estimated extents of direct oxidation of each glucose carbon are, in order: C-l, 28; C-2, 18; C-3, 36; C-3, 4, 47 ; C-4, 52 (estimated from an indirectly derived slope); C-6, 20% (Fig. 2). A weighted average (accounting for absence of data for C-5) provides an overall fraction of about 31% if the calculated configuration of C-4 activity is used and about 28 % if data from [3, 414C]glucose is used directly instead. These estimates corres-
AND
WANG
pond with the only other estimate recorded under directly comparable conditions, of 30 % (Depocas, 1964). Most estimates of prompt oxidation of glucose in infusion experiments have utilized uniformly labeled glucose, and fall between 15 and 25 “/, depending on the extent of fasting and the rate immediately derived from glucose (Shipley et al., 1970; Balasse, 1971). Substrates which enter the glycolytic sequence later than glucose and which ultimately form labeled acetate oxidize in a pattern similar to that of glucose. [6-i4C]Fructose and Na-[2J4C]pyruvate (Fig. 2), and [I J4C]glycerol (not shown) are examples. Oxidation of the first carbon of pyruvate reaches equilibrium quickly, presumably because access to the decarboxylation reaction is immediate, and proceeds thereafter at a relatively constant rate, as do substrates which enter the citric acid cycle directly, such as Na- [1-14C]glutamate (Fig. 2), and Na[ 1J4C]aspartate and Na-acetate labeled in either position (not shown). The rate of conversion of the respective glucose carbons to 14C0, is depressed to varying degrees in intoxicated animals. It is not possible to assign a regular rate of change in each case which can be extrapolated to time zero, as was done with the control experiments. The time courses have a number of characteristics that may be used in evaluation of the metabolic effect of intoxication, however. In intoxicated animals, output of 14C0, was more or less constant from the first equilibrium at about 6 hr through about 4 hr after the infusion of MMH ended, except in the case of [lJ4C]glucose. Apparently, recovery then began, and the rate of oxidation of all labels except C-6 rose rapidly to approach that of untreated animals by the end of glucose infusion. Even though depressed to some degree, carbons 1, 2, and 3 were converted to 14C02 throughout the experiment at a rate greater than the extrapolated direct oxidation rate ; oxidation of [lJ4C]glucose was decreased only slightly.
METABOLIC
TOXICITY
OF METHYL
DISCUSSION
In contrast, C-4 and C-6 were converted to COZ at a much lower rate than the carbons of the other half of the glucose molecule and turned over at a rate lower than the presumed oxidation rate in normal animals. The effect on [6J4C]fructose (Fig. 2) (and on [lJ4C]frustose, not shown) was of the same order as that of [lJ4C]glucose. Oxidation of Na- [I J4C]acetate and Na- [214C]acetate, Na- [ 1-14C]pyruvate, Na- [214C]pyruvate, Na- [ lJ4C]glutamate, and Na[lJ4C]aspartate were unchanged by MMH. The relationship of the respective apparent rates of conversion of the various glucose labels throughout the time course is shown in Table 2. The rate of 14C0, production from each label by intoxicated animals is tabulated as percentage of the rate in control animals at the same point in the time course. In each case, conversion was faster in treated rats early in the experimental period. Additional experiments with [6-L4C]glucase at lower doses show that 0.018 mmol MMH/kg/hr produced a modest inhibition with a similar time course to that of higher doses, and 0.009 mmol/kg/hr had no effect.
Metabolic pathway derangement in MMHtreated, intact animals is rarely studied, and the use of continuously infused labeled substrates in evaluating metabolic activity is also infrequent. As a consequence, the nature of the system should be reviewed to assist in understanding the toxicologic information. Use of a metabolic map may assist in following the discussion. Experiments in which labeled substrate is administered as a single dose have the limitation that a steady state is never achieved; concentration of substrate or intermediates continuously changes in all compartments. This condition may mask changes in rate or extent of reactions which have been induced by toxicity. The value of continuous infusion of substrates was established long ago (Coxon and Robinson, 1959); Shipley et al. (1970).recognized that the approach could dispense with the complex models needed for single dose experiments. Shipley et al. (1970) also suggested, however, that long experiments might compromise the subject and this concern has limited such work to periods of 2-5 hr, usually dealing with regulatory mecha-
TABLE EFFECT
OF MONOMETHYL
HYDRAZINE
231
HYDRAZINE
(MMH)
2 ON
14C0,
FORMATION
FROM
VARIOUS
LABELS
OF [“+C]GLUCOSE
Hours after beginning MMH Begin MMH 4 0
2
Glucose [I-T] [2-‘4CJ [3-Y 1 [3,4-W] 4b [6-W]
Begin glucose 4 4 6
End MMH J 14
16
18
20
22
19 75 108 76 50 40
87 86 107 86
98 93 123 91
45
60
8
10
12
94 70
81 55
75 55
91
88
74
78
loo
86
41 125
75 80
95 110 86 76 69 79
97 100
138
105 130 102
74 55 62
81 74 46
58 47 34
57 30 30
145”
End glucose 4 24
a WO, output, expressed as percentage of output by untreated rats. ’ Calculated as ([(3,4-Y-(3-W)] intox}/{[(3,4-L4C)-(3-‘4C)] control}, where each factor is the respiratory WO, as net counts per minute per 5 min arising from the respective labeled glucose at time (1).
232
DOST,
JOHNSON,
nisms rather than metabolism. We have applied the concept to make a semiquantitative evaluation of the impact of intoxicants on the main avenues of carbohydrate metabolism in intact animals. It is clear from the data that at least 4 to 6 hr of observation is necessary before the impact of intoxication on rates of metabolic activity can be assessed, and 24 hr of observation are needed for full evaluation. The few other investigators who have used continuous infusion of more than traces of glucose have also found a similar delay, but the dose rates and general methods differ so extensively that comparison is difficult (Stretten et al., 1951; Wick et al., 1951; de Freitas and Depocas, 1965; Balasse, 1971; Depocas, 1959; Searle et al., 1956; Bergman, 1963). The biphasic nature of the rate plots for 14C0, output suggests at least two major influences on the time required to achieve a steady state. The most probable governing factors are the rates of release of label from various metabolic pools or sinks. Physical dilution of infused iabeled glucose in the glucose compartments of the body is apparently very fast (Shipley et al., 1967; Searle et al., 1956) and does not contribute to the delay. A comparison by Baker et al. (1959) of extracellular. glucose space compared with total tissue glucose as measured by other investigators (Cori et al., 1953; Orten and Sayers, 1942; Gey, 1956), shows that virtually no glucose exists as such within cells, which means that glucose is diluted only in the extracellular space. Equilibration of metabolic CO, also probably has little influence on the time course. When bi[14C]carbonate is infused continuously, 14C0, output becomes constant as a rate between 90 and 95 % of input, within about an hour (Fig. 2), which is probably equivalent to the distribution and mixing time of metabolic CO,. A similar time course with a more pronounced second phase has been found by Morris and Simpson-Morgan (1963) over a 3-hr period. In theory, randomization of infused glucose to glucose labeled in other positions
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could contribute to the extended period required to reach a steady state, but such effect should be minimal in our studies. Pentose cycle activity is minimal and does not contribute to unequal flow of C-l and C-6 into acetate-derived products. The glucose loading of these experiments should limit gluconeogenic flow of “relabeled” glucose. Available evidence indicates that loading does inhibit reflux of glucose (Parli et al., 1969). In dogs (Cowan et al., 1969) and rabbit liver (Williams et al., 1971), loading of glucose has been shown to maintain C-l/C-6 ratios in 14C02 at about 1, and to prevent endogenous glucose production (Long et al., 1971; Hetenyi and Wrenshall, 1968; von Holt et al., 1961). There are considerable data to indicate that there are extensive metabolic sinks or pools into which carbon enters, no longer as a constituent of glucose. A single dose of labeled glucose disappears from plasma within minutes, transformed into glucose intermediates; labeled CO, does not appear in significant amounts during this early phase (Baker et al., 1959; Baker and Huebotter, 1964). Shipley et al. (1970) have discussed the great capacity of metabolic pools, particularly lipid compartments for glucose carbon prior to oxidation. Whether considered as slowly releasing pools such as fat, or accumulation of more rapidly processed metabolites in the direct glycolytic sequence, when removal is slower than intake the rate of removal becomes rate limiting for release of 14C0 As substrate (or”’ intermediate) builds up there should be a gradual constant increase in the rate of output of 14C0, until some new limiting step is reached or until the entire system is at equilibrium with intake. It seems reasonable to attribute the second phase of gradual and constant increase in rate of 14C0, output to such a series of reactions. The first phase of more rapid change must reflect a period of approach to constant rate among the series of more immediately accessible reactions.
METABOLIC
TOXICITY
Evidence in our experiments supports the contention that the equilibration periods are functions of the time required for carbons to complete reaction sequences. [6-r4C]Fructose and Na- [2J4C]pyruvate which label acetate, require the same period of equilibration as does glucose (Fig. 2), but Na-[1-*4C]pyruvate, which is decarboxylated before any diversion to nonglucose products can occur, is converted very rapidly, and has onIy a limited slope in the second phase. Since Na[ 1J4C]glutamate and Na- [ 1-14C]aspartate are known to enter the citric acid cycle readily and they are immediately oxidized, it may be implied that the cycle moves quickly and is not linked with a metabolic sink for carbon. In view of the respective behavior of acetatelabeling and pyruvate-l-labeling precursors it is fair to assume that flux of acetate into fatty acid synthesis and oxidation temporarily disposes of a large amount of glucose carbon. Oxidation of glucose, fructose, and [2J4C]pyruvate are all similarly delayed, but oxidation of [I-r4C]pyruvate is rapid. The difference between the relatively slow turnover of C-3 of glucose, which eventually becomes carbon-l of pyruvate, and the rapid oxidation of exogenous [lJ4C]pyruvate can be explained only by the time required for biochemical equilibration of glucose at other points in the sequence. The times required for overall isotopic equilibration as judged by 14C0, production are more or less constant at infusion rates from traces of [r4C]glucose up to 500 mg/hr. Furthermore, if we infuse 150 mg unlabeled glucose/hr to rats under the conditions of experiments reported here and after 6 hr add a trace label to the continued glucose infusion, the expected 4- to 6-hr elapse before the first equilibration phase as represented by 14C0, output is completed. A single priming dose of labeled material superimposed on the system in an effort to establish an earlier steady state will influence all compartments unequally for about the same period and provides no advantage in continuous infusion experiments.
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Changes in 14C02 output resulting from acute intoxication during infusion are not subject to the same delays because all reactions have reached maximum rates, dependent either on availability of substrate or reaction capacity. Animals with previously established toxicity are subject to the same kinds of time course, although specific rates within the system may be changed, and the most useful information is the rate data obtained after the first phase is complete. Acute non lethal fluoride intoxication, for example, which apparently blocks glucose entry to cells while causing no obstruction in main pathways, results in an almost immediate drop in 14C0, production from all [14C]glucose labels (Dost et al., 1977). Interpretation of Changes Resulting from Intoxication In the process of interpreting observed toxic effects it is first necessary to establish whether the change is in the direct oxidation of glucose (glycolysis, pentose cycle, and the citric acid cycle) or on biosynthetic pathways into which carbon may move for later return and oxidation. If MMH primarily influenced movement of carbon into or out of reaction compartments not in the direct line of metabolism, a much greater symmetry of effect on the two halves of the glucose molecule should be evident. We should also see an appreciable difference between oxidation rates for C-3 when compared with C-2, and for C-4 compared with C-6. The sharp singular decrease in C-6 oxidation and the decrease of C-4 and C-6 oxidation below the level of estimated direct metabolism indicates oxidation of the C-4, 5, 6 half of the molecuIe is selectively obstructed, which can only happen in a main pathway, probably glycolysis. The rate of conversion of continuously infused [6J4C]glucose to 14C0, will represent, within limits, the total glycolytic flux because pentose cycle activity is normally slight and the glucuronic acid pathway is negligible. A small additional fraction of the 14C0, from
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C-6 will arise from glucose cycled through the pentose pathway; of every three glucose molecules entering that pathway, one terminal 3-carbon fragment (C-4, 5, 6) will enter triose metabolism as glyceraldehyde-3-phosphate. MMH caused a marked decrease in production of 14C0, from [6J4C]glucose and a similar apparent decrease in conversion of carbon-4. These changes indicate glycolytic impairment, since the 4,5,6-carbon fragment must remain intact through either glycolysis or the pentose cycle until carbon 4 is lost in decarboxylation of pyruvate. The rate of conversion of [6-r4C]glucose under those conditions will then represent the amount of intact glucose passing through glycolysis, including that reformed during pentose cycle activity, and less that arising from glyceraldehyde-3-phosphate which was formed from a terminal 3-carbon unit in the pentose pathway. Differentiation of these two components is difficult if not impossible to precisely measure in an intact animal system. Carbons 1, 2, and 3 of glucose may be assumed to traverse glycolysis proper to the same extent as carbon 6. The extent of C-l oxidation in excess of C-6 oxidation may be taken as an indication of increased activity of the pentose pathway. A similar effect of acetyl phenylhydrazine on glycolysis has been observed in erythrocytes (Szeinberg and Marks, 1961; Kosower et al., 1963; Mills and Buell, 1965). Having established that glycolysis was impaired, it was necessary to learn where in this rather extensive sequence the defect might be. (The possibility of interference in the citric acid cycle was considered unlikely since oxidation of Na- [ 1J4C]pyruvate and Na- [2J4C]pyruvate, and Na [ lJ4C]aspartate, and Na- [ 1J4C]glutamate were all unaffected by MMH.) The next step was to determine whether the effect was on hexose or triose metabolism. Fructose is the best substrate for this problem because it enters glycolysis rapidly in normal
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animals as glyceraldehyde phosphate and dihydroxyacetone phosphate. The limited decrease in [6-14C]fructose catabolism under MMH intoxication indicates that consumption of triose was not seriously slowed, and directed attention to hexose catabolism. (Use of [lJ4C]fructose provided identical information but is not reported.) On the basis of the considerably greater conversion to 14C0, of labeled C-2 and C-3 than C-6, some aspect of hexose metabolism is clearly capable of proceeding. This can only occur if fructose 6-phosphate formed in the pentose cycle was readily transformed to glucose 6-phosphate in the bidirectional phosphoglucose isomerase reaction. Since the fructaldolase conversion of fructose 1,6-phosphate to trioses was not impaired it is reasonable to assume that fructose 1,6phosphate metabolism to trioses was also unaffected since the same enzyme is required. The remaining step and therefore the logical site where inhibition must occur is the key regulatory phosphofructokinase reaction.
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