Phosphate-dependent glutaminase of small intestine: Localization and role in intestinal glutamine metabolism

ARCHIVES

OF

BIOCHEMISTRY

AND

BIOPHYSICS

182, 506-517 (1977)

Phosphate-Dependent Glutaminase of Small Intestine: Localization and Role in Intestinal Glutamine Metabolism’ LAWRENCE Nutritional

Biochemistry

M. PINKUS?

Section, Metabolism

AND

HERBERT

Laboratory ofNutrition and Digestive Diseases,

G. WINDMUELLER

and Endocrinology, Bethesda, Maryland

National 20014

Institute

ofArthritis,

Received January 25, 1977 An important metabolic function for glutamine in rat small intestine is suggested by the large uptake from blood and metabolism to account for about 30% of CO* produced [Windmueller, H. G., and Spaeth, A. E, J. Biol. Chem. (1974) 249, 5070-50791. The activities of several glutamine-degrading enzymes in intestine were therefore examined. A highly active phosphate-dependent glutaminase was found in the mucosal epithelium of rata, dogs, cats, hamsters, rabbits, and monkeys; activity was lower in guinea pigs and chickens. The net uptake of glutamine by intestine was greatest in those animals with the highest blood glutamine concentration and tissue glutaminase activity. Specitic activity was similar in mucosa of rat duodenum, jejunum, and ileum, but diminished in stomach, cecum, and colon. Phosphate-dependent glutaminase was found in both villus and crypt cells and was associated with mitochondria. The enzyme appears similar in many respects to phosphate-dependent glutaminase of kidney and brain. However, unlike the kidney enzyme, its activity was not induced during acidosis. The glutamine analogs, L-2-amino-4-oxo-5-chloropentanoic acid (chloroketone) and 6-diazo-5-oxo-L-norleucine (DON) were effective irreversible inhibitors in vitro and in vivo. Glutamine protected against inhibition. Chloroketone administered luminally into a rat jejunal segment in situ resulted in 50% inhibition of glutaminase activity and totally inhibited the hydrolysis of glutamine subsequently absorbed from the lumen. The hydrolysis of glutamine taken up from arterial blood was not affected. However, DON administered arterially inhibited glutaminase almost completely and inhibited the hydrolysis of both luminal and arterial glutamine. The dam indicate that metabolism of glutamine to glutamate occurs in mitochondria of both villus and crypt cells and that phosphatedependent glutaminase is the major glutamine-utilizing enzyme of small intestine.

Recent studies have described the uptake and metabolism of plasma glutamine by the small intestine in rats and in a number of other animal species (1, 21, including man (3). From arteriovenous difference measurements in rats, intestine is the only tissue which exhibits appreciable net glutamine uptake (l), about 170 nmol/ 1 Portions of this work were presented at the 67th annual meeting of the American Society of Biological Chemists in San Francisco, California, June 610, 1976, and at the 10th International Congress of Biochemistry in Hamburg, West Germany, July 25 31, 1976. 2 Present address: Department of Gastroenterology, Nassau County Medical Center, East Meadow, New York 11554. Copyright 0 1977 by Academic Press, Inc. All rights of reproduction in any form reserved.

min/g of tissue (2). About one-third of the total plasma glutamine pool is extracted in a single pass through the intestinal vasculature. Less than 15% of the L-HI14C]glutamine taken up by rat intestine from arterial blood (1) or from the intestinal lumen (2) is incorporated into tissue protein; the remaining glutamine carbon is rapidly metabolized to carbon dioxide (63%), organic acids (20%), and other amino acids (12%). Plasma glutamine is the source for about 30% of the total carbon dioxide released by the gut in vivo and thus serves as an important substrate in the respiratory metabolism of intestinal tissue. In further studies of the pathway of glu506 ISSN 0003-9861

GLUTAMINASE

IN

tamine metabolism in intestine, we have now identified phosphate-dependent glutaminase (L-glutamine amidohydrolase, EC 3.5.1.2) as the major glutamine-utilizing enzyme and present a detailed study of its activity. MATERIALS

AND

METHODS

Materials. Except where indicated, OsborneMendel adult male rats fed NIH-07 open formula stock ration were used. Other animals were obtained from National Institutes of Health animal colonies and chickens were obtained from local farms. Ammonia-free glutamate dehydrogenase (EC 1.4.1.2) was obtained from Sigma and rH]thymidine (23-27 Ci/mol) from Amersham/ Searle. All chemicals were of the highest grade commercially available. L-2-Amino-4-oxo-5chloropentanoic acid (chloroketone4) was synthesized as described previously (41, and 6-diazo-5-oxo-L-norleucine (DON) was obtained from the Drug Development Branch, National Cancer Institute. Tissue preparations. Mucosa was obtained from excised segments of intestine after rinsing the lumen thoroughly with phosphate-buffered saline. Adhering mesentery was trimmed, the segments were opened longitudinally, and after additional mucus was removed by blotting with moist tissue paper, mucosa was freed from the underlying muscularis by scraping with a glass slide on an ice-cold surface. Homogenates were prepared by disruption of mucosal scrapings in 10 vol of 125 mM potassium phosphate, 330 mM sucrose, 2 mM dithiothreitol (pH 7.6) (homogenization buffer) in a Waring Blendor at O6°C for 45 s. Alternately, 20 strokes of a motordriven Teflon-glass homogenizer were used. To prepare homogenates of whole intestine, a 10-s treatment with an Ultra-Turrax homogenizer was found to be most effective. As noted by Stern (51, the presence of a mercaptan disrupts mucus which can interfere with subsequent differential centrifugation and pipetting procedures. To prepare mitochondria, homogenates of intestinal mucosa prepared in a Teflon-glass homogenizer were first centrifuged for 10 min at 800 g to remove nuclei and cell debris. After aspiration of floating fatty material, the mitochondria were sedimented by centrifugation for 10 min at 15,OOOg. The mitochondrial pellet was twice washed by resuspension in homogenization buffer without dithiothreitol

4 Abbreviations used: chloroketone, L-2-amino-4oxo-5-chloropentanoic acid; DON, 6-diazo-5-oxo-Lnorleucine; Hepes, N-2-hydroxyethylpiperazine-N’2-ethanesulfonic acid.

INTESTINAL

MUCOSA

507

in a glass Dounce homogenizer (type A) followed by centrifugation. A 105,OOOg supernatant, essentially free of glutaminase, was obtained after homogenization of intestine in a solution containing 100 mM Tris-HCl, 50 mM KCl, 1 mM MgCl,, 0.5 mM EDTA, 2 mM dithiothreitol, and 30% glycerol (pH 7.6). This was used as a source of amidotransferases and glutamine transaminase. Rat mucosal villus and crypt cells were separated by the method of Weiser (6). The cell fractions washed from the lumen were centrifuged at 500 g for 3 min, rinsed twice with phosphate-buffered saline at 4”C, and the volume of packed cells was noted. Villus cells were released first, and crypt cells appeared in later washings. The cells prepared in this way were virtually free of mucus. Cell homogenates were prepared in 8 vol of 125 mM potassium phosphate, 330 mM sucrose (pH 7.6) with 30 strokes of a Dounce homogenizer (type B) and were centrifuged for 10 min at 15,000 g. The resulting pellets (glutaminase) and supernatant fractions (thymidine kinase and alkaline phosphatase) were used for determination of enzyme activities. Determination of enzyme actiuities. All assays were carried out at 37°C. Phosphate-dependent glutaminase activity was determined, except where noted, by measuring glutamate formation with glutamate dehydrogenase (see below). Membrane preparations, as indicated in each experiment, were incubated in 135-150 mM potassium phosphate, 16.7 mM glutamine (pH 8.1); the final volume was 0.3 ml. The reaction was terminated by adding trichloroacetic acid to a final concentration of 5%. Protein was removed by centrifugation and aliquots of the supernatant fraction were withdrawn for determination of glutamate. Determinations were corrected using incubation mixtures to which enzyme was added after trichloroacetic acid. Phosphate-independent glutaminase was assayed as described by Tate and Meister (7) or Curthoys and Kuhlenschmidt (8). Homogenates prepared in 50 mM Tris-HCl or 50 mM Hepes containing 330 mM sucrose (pH 7.5) were used as the source of the enzyme. Reactions were stopped with trichloroacetic acid and the glutamate formed was determined. y-Glutamyltranspeptidase (EC 2.3.2.2) was assayed with L-y-glutamyl-p-nitroanilide as described by Orlowski and Meister (9) with the addition of glycylglycine as acceptor (10). Cytochrome oxidase (EC 1.9.3.1) was assayed as described by Smith (ll), and NADPH-cytochrome c reductase (EC 1.6.99.1) by the procedure of Williams and Kamin (12). Published conditions were employed for the assays of glutamine transaminase (EC 2.6.1.15) using glyoxylate as acceptor (131, carbamoyl phosphate synthase (EC 2.7.2.9) (14), phosphoribosyl pyrophosphate amidotransferase (EC 2.4.2.14) (15), cytidine triphosphate synthetase (EC 6.3.4.2) (16), and fruc-

PINKUS

508

AND WINDMUELLER

tose-6-phosphate aminotransferase (EC 2.6.1.16 = EC 5.3.1.19) (17); activities, however, were monitored through the disappearance of glutamine or the formation of glutamate dependent upon acceptors specific for each reaction. Thymidine kinase (EC 2.7.1.21 = EC 2.7.1.75) activity was determined by incubating aliquots of a 15,000g supernatant at 37°C in a solution containing 50 mM Tris-HCl, 5 mM ATP (18), 5 mM MgCl,, and 4 PM [3H]thymidine (pH 8.0); the final volume was 0.1 ml. Aliquots were withdrawn at intervals for 15 min and the phosphorylated products were collected on Whatman DE-81 filters and quantitated as described by Breitman (19). Alkaline phosphatase (EC 3.1.3.1) was determined as described by Weiser (6). O&r procedures. Glutamate was determined with glutamate dehydrogenase (20). Aliquots (50-75 ~1) of a deproteinized assay mixture containing no more than 0.25 pmol of glutamate were added to 0.75 ml of 0.5 M glycine and 0.4 M hydrazine hydrate (pH 9.2). Following the addition of 0.075 ml of a solution containing 50 mM sodium phosphate, 50% glycerol, 2 pmol of NAD, and 24 units of glutamate dehydrogenase (pH 7.3), the mixture was incubated for 35 min at 22-24°C and the increase in absorbance at 340 nm was measured. Recovery of glutamate was 90-100%. Glutamine and ammonia were enzymatitally determined as previously described (1). Protein was determined by the method of Lowry et al. (21). RESULTS

Glutamine-Utilizing Intestine

Enzymes

of Small

To define the major pathway for glutamine metabolism in gut we measured the activities of several glutamine-degrading enzymes and obtained additional data from a survey of scattered reports in the literature. An active phosphate-dependent glutaminase was found in homogenates of mucosal scrapings (specific activity, 3-6 pmol/h/mg of protein). Phosphate-independent glutaminase activity was extremely low, about 1% that of phosphatedependent activity (see Discussion). A 105,000 g supernatant fraction of an intestinal homogenate (see Methods) was free of glutaminase activity and therefore was used to survey soluble glutaminedegrading enzymes, i.e., glutamine transaminase and amidotransferases. Glutamine transaminase was detectable (0.1 km01 of glutamine lost/h/mg of protein) while several amidotransferases (carbamoyl phosphate synthetase, phosphori-

bosyl pyrophosphate amidotransferase, cytidine triphosphate synthetase, and fructose-6-phosphate aminotransferase), monitored through glutamate formation, were undetectable, indicating specific activities of less than 0.07 pmol/h/mg of protein. Our results are in accord with previous reports which indicated the presence of a number of glutamine-degrading enzymes in rat small intestine. In addition to phosphate-dependent glutaminase (1.8-12 pmol/h/mg of protein) (22-25), low activity had been noted for glutamine transaminase (261, carbamoyl phosphate synthetase (271, phosphoribosyl pyrophosphate amidotransferase (281, and fructoseS-phosphate aminotransferase (29-31); activities reported were 0.16, 0.07, 0.07, and 0.0120.036 pmol/h/mg of protein, respectively.5 Although subject to the uncertainties associated with determination of enzyme activities in crude preparations, the results indicate that phosphate-dependent glutaminase is by far the most active enzyme of glutamine metabolism in gut and is the only one with sufficient activity to account for the in uiuo rate of glutamine metabolism. We therefore sought to localize and characterize this activity. Intestinal Glutaminase Activity ent Species

in Differ-

Phosphate-dependent glutaminase activity was detected in the mucosa of small intestine in all the animals surveyed (Table I). The specific activity was generally similar in proximal and distal small intestine; however, higher activity was found in the jejunum of the dog and in the ileum of the cat. The enzyme specific activity in most animals was similar to that found in the rat. Exceptions were the chicken and guinea pig, where activity was lower. It is noteworthy that in these two species the arterial glutamine concentration is also lower,6 and, contrary to what is found in 5 Where activity was expressed per gram of tissue, activity was recalculated per milligram of protein assuming 100 mg of protein/g of tissue, wet weight. 6 The low glutamine concentration in guinea pig blood cannot be ascribed to the circulating asparaginase since this enzyme is reportedly devoid of glutaminase activity (32).

GLUTAMINASE

IN

INTESTINAL

TABLE PHOSPHATE-DEPENDENT

GLUTAMINASE

Animal

Tissue

Dog (3) Rat (12) Hamster (3) Cat (2) Mouse (4) Monkey (1) Rabbit (2) Guinea pig (3) Chicken (2)

ACTIVITY

MUCOSA

(pmol/

Jejunum

Ileum

5.1 4.8 3.6 1.6 4.9 4.7 4.0 1.2 0.4

2.8 5.0 4.3 4.8 5.1 4.9 5.3 1.4 0.3

FROM

DIFFERENT

Arterial glutamine concentration* (pmol/liter of plasma)

ANIMAL

SPECIES

Arteriovenous difference for glutamine across intestine (V-A)b (“mo&t$ of

723 655 741 602 -

-200 - 194 -153 -129 -

463 433 247 330

-148 -40 +14 +53

animals indicated. Intestine was obtained from freshly killed the ligament of Trietz and the ileocecal valve. The proximal portion as ileum. Mucosal scrapings were used as the source taken from Windmueller and Spaeth (1). Blood sampling described (1) were used to obtain results with chickens, mean body weights of 290 g.

the other animals studied, the arteriovenous difference for glutamine concentration across the gut is positive, indicating a net output rather than a net uptake of circulating glutamine by the intestine (Table I). Among these animals, there was an apparent positive correlation between the arterial glutamine concentration, the extent of net uptake of circulating glutamine, and the tissue glutaminase activity (Table I). Gastrointestinal

I

OF INTESTINAL

glutaminase activity” h/mg of protein)

cLData are mean values for the number of animals and divided at a point midway between portion was regarded as jejunum and the distal of enzyme. * Data for all species except chicken were analytical methods similar to those previously male and one female of Leghorn strain with

509

MUCOSA

Distribution

The glutaminase distribution in the gastrointestinal tract of the rat was determined in more detail. Activity was distributed fairly uniformly along the entire length of the small intestine; the specific activity was similar in duodenum, jejunum, and ileum. Lower specific activities were observed in the stomach, cecum, and colon (Table II). When the mucosa and muscularis of rat small intestine were separated by scraping, the mucosa, which comprised about 60% of the total intestine weight, contained over 90% of the total glutaminase activity. This value approached 96% in epithelial cells freed from the muscularis by the Weiser procedure (6). We conclude that virtually ali the glutaminase in rat

TABLE

and three

II

DISTRIBUTION OF PHOSPHATE-DEPENDENT GLUTAMINASE ACTIVITY IN RAT GASTROINTESTINAL MUCOSA Tissue”

Specific activity (cLmol/h/ mg of protein)

Stomach Duodenum Jejunum (upper) Jejunum (lower) Ileum (upper) Ileum (lower) Cecum Colon

0.5 3.6 4.2 3.9 3.6 3.5 1.0 0.8

Total a Mucosal scrapings enzyme. Data represent ments.

Percentage of total activity

2 9 17 19 21 21 6 5 100

were used as the source of the average of two experi-

small intestine is confined to the epithelial cells. Mucosal epithelial cells are produced in the crypts and undergo morphologic and enzymatic changes as they migrate to the tips of the villi (33-35). To determine if glutaminase is localized in a particular class of epithelial cells, the villus and crypt cells were separated by successive washes of the intestinal lumen (Fig. 1).

510

PINKUS Villus

0'

AND

Crypt

I I I I 20 40 al so % Total Packed Cell Volume

I 1w

FIG. 1. Distribution of glutaminase activity in villus and crypt cells of intestinal mucosa. Epithelial cells were sequentially eluted from a 2.5-g segment of jejunum by the method of Weiser (6). About 3.0 ml of packed cells were obtained from the segment. One milliliter of packed cells contained approximately 80 mg of protein.

Villus cells appeared in early washes and were identified by high alkaline phosphatase activity while crypt cells appeared in later washes and were identified by high activity for thymidine kinase. The specific activity of glutaminase, expressed per milligram of protein, was approximately the same in all cell fractions. Mitochondrial

WINDMUELLER

jected to isopycnic centrifugation in a sucrose gradient (Fig. 2), a single peak corresponding to the density of mitochondria (38) was observed containing both glutaminase and cytochrome oxidase. Significant quantities of phosphate-dependent glutaminase activity can be extracted from lyophilized kidney mitochondria with hypotonic phosphate-borate buffer (39, 40). Preliminary studies indicate that some intestinal glutaminase activity can also be solubilized in this way. Therefore, further purification of the enzyme should be possible. Properties

of the Enzyme

Phosphate-dependent glutaminase was studied using mitochondria from rat intestinal mucosa as the enzyme source. In the phosphate-containing homogenization buffer, the enzyme retained over 90% of its activity after 30 min at 37°C and 6070% after 24 h at 4°C. Preparations exhibited a broad pH optimum between pH 7.6 and pH 9.2, but activity fell to near zero below pH 6.0 or above pH 10.2. At pH 8.1 the apparent K, for glutamine was 2.2 mM. Enzymatic activity was retained in preparations of lyophilized mitochondria after rehydration in homogenization

Localization

Differential centrifugation of a homogenate of rat small intestinal mucosa gave the following fractions: (a) nuclei and unbroken cells (500 g, 10 min); (b) mitochondria (15,000 g, 10 min); (c) microsomes (105,000 g, 90 min); (d) 105,000 g supernatant. The fractions contained approximately 30-35, 57-66, 3-6, and l-2% of the total phosphate-dependent glutaminase activity, respectively. Glutaminase and the mitochondrial enzyme, cytochrome oxidase, were three- to fourfold enriched in the 15,000 g pellet compared to the total homogenate. Specific activity was greatly decreased in the microsomes (enriched in NADPH-cytochrome c reductase). Purified brush borders (36) and nuclei (37) were virtually devoid of glutaminase activity. When the mitochondrial fraction was sub-

FIG. 2. Sucrose density gradient centrifugation of intestinal mitochondria. A washed, resuspended 15,OOOg pellet of mitochondria (1 ml, 10 mg protein) was layered over a 16-ml gradient of 30-60% sucrose containing 125 mM potassium phosphate (pH 7.4). Centrifugation was carried out at 105,000 g for 12 h in a Beckman SW 27 rotor at 4°C. Fractions (0.6 ml) were collected and assayed for phosphate-dependent glutaminase and cytochrome oxidase. Sucrose density was determined by refractometry.

GLUTAMINASE

IN

buffer, and the apparent K, for glutamine was unchanged. This suggests that disruption of the mitochondrial membrane does not alter the apparent affinity of the enzyme for substrate and that under our assay conditions the permeability of the mitochondrial membrane to glutamine is not rate limiting for enzyme activity. The effectiveness of phosphate and other anions as activators was tested (Table III). Mitochondria prepared in 50 mM Tris, 330 mM sucrose (pH 7.6) exhibited virtually no glutaminase activity upon incubation with glutamine in Tris buffer (pH 8.0) and were thus a suitable source of the enzyme. Although phosphate was the best activator, other compounds, including glucose-6-phosphate and ammonium sulfate, were almost as effective. Plots of activity versus concentration for all three compounds were hyperbolic with 22 mM. apparent Ko., of approximately The enzyme displayed more complex activation curves with maleate, pyrophosphate, and ammonium bicarbonate, but apparent K,,, values were also in the range of lo-30 mM. Activation by glucose6-phosphate and pyrophosphate could not TABLE ACTIVATION GLUTAMINASE Addition

III

OF PHOSPHATE-DEPENDENT BY VARIOUS COMPOUNDS”

to assay

mixture

Phosphate Ammonium sulfate Glucose-6-phosphate Sulfate Maleate Ammonium bicarbonate Oxalate Pyrophosphate Citrate Potassium Arsenate None

Percentage tivity

of ac-

100 100 95 76 67 62 59 53 43 34 0

a Aliquots of a solution containing washed mitochondria from rat intestinal mucosa were incubated at 37°C for 20 min in an assay mix containing 25 rnM Tris-HCl (pH &X0), 16.7 mM glutamine, and a 67 rnM concentration of the indicated anion or organic acid (added as the sodium salt unless otherwise indicated and adjusted to pH 8.0 with NaOH). The glutamate formed was determined.

INTESTINAL

511

MUCOSA

be attributed to release of inorganic phosphate during assay because (a) only very low levels of alkaline phosphatase activity were found in the mitochondrial preparations under the assay conditions employed (< 0.2 pmol of p-nitrophenol released/h/mg of protein), and (b) glutaminase activity was linear with time for these activators. Succinate, phthalate, ammonium formate, ammonium acetate, EDTA, and AMP activated weakly (<15%), while formate, lactate, cr-ketoglutarate, pyruvate, fumarate, glycine, Laspartate, L-serine, sodium carbonate, ammonium chloride, Hepes, and cyclic AMP did not appear to activate. Intestinal homogenates or mitochondria from rat intestine did not release ammonia from n-glutamine or L-asparagine; rat intestine in uiuo also failed to hydrolyze Lasparagine (41). Thus L-glutamine appears to be the principal substrate. Mitochondria from dog intestine displayed weak asparaginase activity equal to about 10% of the glutaminase. Inhibition

Studies

in vitro

Glutaminase was inactivated (>99%) upon preincubation of rat intestinal mitochondria at 37°C for 30 min in 125 mM potassium phosphate (pH 7.5) with the sulfhydryl inhibitor p-hydroxymercuribenzoate (0.5 111~). DON and chloroketone, glutamine analogs which are highly specific irreversible inhibitors of glutamine-utilizing enzymes (42), also inhibited the enzyme (Table IV). Virtually complete inhibition could be obtained with either compound, although at lower concentrations DON was more effective. Glutamine offered significant protection. Reduced activity was also observed in mucosal scrapings obtained after intact jejunal segments were incubated in vitro at 37°C for 30 min with phosphate-buffered saline containing either 5 mu DON or 20 mM chloroketone in the lumen. Glutamine (100 mM) also protected about 50-60% of the activity under these conditions. The protection by glutamine suggests that the analogs inhibit by reaction with glutamine binding site(s) of the enzyme.

512

PINKUS TABLE

INHIBITION GLUTAMINASE Inhibitor

AND

WINDMUELLER

IV

Inhibition

OF PHOSPHATE-DEPENDENT BY CHLOROKETONE AND DON”

concentration b-f)

0 2.5 2.5 (+ 100 mM glutamine) 5 10 10 (+ 100 mM glutamine) 20

Chloroketone

(100) 10 93 4 3 1

0.w 41 22 14 91 6

’ Intestinal mitochondria were incubated for 30 min at 37°C in a solution containing 125 mM potassium phosphate (pH ‘7.6) and the indicated concentration of inhibitor or inhibitor plus glutamine. The mixtures were then cooled to O”C, dialyzed for 2 h against 1000 vol of 125 mM potassium phosphate, 330 mM sucrose (pH 7.6), and assayed for glutaminase. TABLE INHIBITION Line

Jejunal

segment

1 2

Control Control

3 4

Experimental Experimental

5

Experimental

OF GLUTAMINE Addition

Earle’s Earle’s mine None Earle’s ketone’ Earle’s mine

HYDROLYSIS to lumen

V BY CHLOROKETONE Duration (min)

Net

Output

Glutamine, plasma

IN

Vrvoa (nmol/min/g

of tissueY

Ammonia Plasma

Perfusate

Total

gluta-

25 30

-220 72

73 136

59 125

132 261

+ 20 mM chloro-

25 40

-288 -258

128 154

-

128 154

103

43

146

salts (perfused) salts + 6 mM (perfused) salts

in vivo

The effects of the glutamine analogs were also tested on isolated jejunal segments in vivo (2). Normal rates of uptake and hydrolysis of arterial and luminal glutamine by a typical untreated segment are shown in Table V (control). Glutamine from arterial blood was utilized at a net rate of 220 nmol/min/g of tissue (line l), represented as a minus output. Ammonia output, resulting from glutamine hydrolysis, was measured in venous plasma and in the luminal per&sate and totalled 132 nmol/min/g. When 6 mM glutamine was added to the luminal perfusate (line 2) the glutamine concentration increased in venous blood, resulting in a shift from net glutamine uptake to net output (from -220 to + 72 nmobminlg); ammonia output also increased, from 132 to 261 nmol/min/

Glutaminase activity (percentage of control) DON

Studies

salts + 6 mM (perfused)

gluta-

30

133

a The small intestine of an anesthetized rat was exteriorized through an abdominal incision into a 37°C tissue bath containing Earle’s balanced salt solution with 5.6 mM glucose and 0.6 mM glutamine. The intestinal lumen at each end of a 2- to 5-cm segment ofjejunum was cannulated, using ligatures around the intestine to isolate the segment from the rest of the bowel. All venous blood was collected from a cannula in the single vein draining the segment; the arterial blood supply to the segment remained intact. Arterial blood was sampled periodically from a cannula in the abdominal aorta. Replacement blood from donor rats was transfused continuously into a saphenous vein at a rate sufficient to maintain arterial blood pressure. Results are from one control and one experimental segment in separate animals. See Ref. (2) for additional experimental details regarding the intestinal preparation. b Net output into plasma was determined from the arteriovenous concentration difference, determined three to six times during each interval, multiplied times the flow rate of blood through the segment (2). Net output into the luminal per&ate was determined from the inflow-outflow concentration difference multiplied by the perfusate flow rate (0.5 ml/min). Glutamine and ammonia were determined in deproteinized samples as described previously (1). All intestinal uptake of blood glutamine is from the plasma (21. Ammonia, however, equilibrates rapidly between plasma and erythrocytes; therefore the rates given for ammonia output into plasma are equivalent to approximately 65% of the total ammonia output into the blood. Data are the means of three to six values. Standard error values were less than 10% of means. c Chloroketone (0.3 ml) was infused into the segment lumen at the start of this interval and left in place for 40 min.

GLUTAMINASE

IN

g, reflecting the cleavage of luminal glutamine during transport (2). Before chloroketone treatment, the experimental segment took up arterial glutamine and released ammonia into venous plasma at a normal rate (line 3). Infusion of 20 mM chloroketone into the lumen had little effect on either the uptake of arterial glutamine or the release of ammonia (line 41, which was all recovered in the venous blood because the lumen was not perfused. Forty minutes after chloroketone infusion, the lumen was perfused with 6 mM glutamine, which was extensively transported from lumen to blood, as shown by the increase in glutamine output from -258 to +133 nmol/min/g (line 5). However, in contrast to results with the control segment, no increase in ammonia output was seen. Thus glutamine was transported across the intestine without measurable hydrolysis. Tissue analysis following the chloroketone treatment showed that the glutaminase activity of the segment was about 50% inhibited compared with adjacent untreated areas of intestine. Apparently, that portion of the tissue glutaminase localized along the path of absorption, presumably in the villus cells, was inhibited. Absorbed glutamine, following the same path across the tissue as the previously absorbed inhibitor, thus encountered only inhibited enzyme. Another portion of the tissue glutaminase was apparently inaccessible to the chloroketone and was responsible for the continued hydrolysis of glutamine taken up from the blood. In another experiment with a similar intestinal preparation, a 240 mM solution of DON was infused continuously into the arterial blood supply to the segment for 30 min at 5% of the rate of blood flow. Thus the final blood DON concentration was about 12 mM. Since venous blood from the segment was collected, DON did not enter the general circulation and only the experimental gut segment was exposed to the inhibitor. During the DON treatment and over the ensuing 20 min the rate of glutamine uptake from blood gradually diminished in the segment from 251 to 57 nmol/ min/g of tissue; ammonia output was also

INTESTINAL

MUCOSA

513

reduced, from 160 to 75 nmol/min/g of tissue. As in the chloroketone-treated segment, glutamine subsequently perfused through the lumen appeared in the venous blood but did not increase ammonia output. At the end of the experiment glutaminase activity in the segment was inhibited about 97%, indicating that DON can be taken up from blood by all epithelial cells and can gain access to all of the mitochondrial enzyme. Intracellular accumulation of glutamine could explain the residual net uptake of arterial glutamine observed following the DON treatment. A six-fold increase in the mucosal concentration of glutamine has, in fact, been observed in the jejunum of rats following the intravenous administration of DON (1). After both of the in uiuo inhibition studies described above, the mucosal tissue of the treated segments was visibly edematous. In addition to glutaminase, several of the amidotransferases were undoubtedly inhibited by the glutamine analogs (43) so the morphological changes were not necessarily due to impaired glutaminase activity alone. The results, however, suggest that depriving the intestine of its glutamine-utilizing reactions can rapidly produce adverse effects. Effect of Age and Altered States

Physiological

As previously noted, glutaminase activity is increased about fourfold in the rat kidney during acidosis (44, 45). However, conditions that inchased activity in the kidney did not increase the activity in intestine (Table VI). Alkalosis did not appreciably affect the activity in kidney (45) or intestine. No change was observed in the specific activity of glutaminase in the intestinal mucosa when the drinking water of young adult rats was supplemented with 275 mu glutamine for up to 7 days. This suggests that enzyme activity is not induced in response to a greater intake of dietary glutamine. The specific activity of the enzyme was also not changed significantly following an overnight fast. In a study of the temporal development of the enzyme, adult levels of glutaminase

514

PINKUS

EFFECT OF DEPENDENT

AND WINDMUELLER

TABLE VI ACIDOSIS AND ALKALOSIS

Condition

Normal Acidosis Alkalosis

ON PHOSPHATEGLUTAMINASE ACTIVITY OF INTESTINE AND KIDNEYS

Glutaminase activity (pmol/h/ mg of protein) Intestine 3.1 3.0 2.8

Kidney 5.1 20.7 4.9

’ Acidosis and alkalosis were induced by the addition of 0.28 M NH&I or 0.28 M NaHCO,, respectively, to the drinking water of rats for 7 days. Homogenates prepared from whole intestine and whole kidney were used as the enzyme source. Data are means of two experiments.

(pmol/h/mg of protein) were found in the small intestine of fetal rats immediately prior to birth and up to 5 days of age. Activity then gradually increased about twofold by days 12-15 before returning to the adult level at 20 days of age. DISCUSSION

In the present studies we have identified an active phosphate-dependent glutaminase localized in the mitochondria of mucosal villus and crypt cells in rat intestine. The activity is virtually absent from the muscularis obtained by scraping off the mucosa, or from the residue remaining after selective elution of epithelial cells. This indicates that metabolism of glutamine by glutaminase is restricted to the mucosal epithelial cells and correlates well with the high rate of glutamine metabolism in mucosa relative to muscular-is previously observed in viuo (1). The potential of intestinal phosphatedependent glutaminase for glutamine metabolism can be estimated from the specific activity of the enzyme (approximately 300 pmol/h/g of intestine), the observed concentration of glutamine in the mucosa (0.25 mu in the cell water) (l), and the apparent K, of the glutaminase (2.2 mM). When sufficient concentrations of activators are present in Go, the concentration of glutamine in mitochondria may be rate limiting in the action of glutaminase. If the steady-state concentration of glutamine in intestinal mitochondria is as-

sumed to be 0.25 mM, the same as in the cell water, then the effective rate of glutaminase activity calculated from the Michaelis-Menten equation would be 30 pmollhlg of tissue. The highest observed rate of glutamine utilization by rat intestine, when glutamine is being taken up from both the lumen and the blood, is 20 pmol/h/g of tissue (2). The action of glutaminase alone seems sufficient, therefore, to account for the highest observed rate of glutamine utilization. The decreased levels of intestinal phosphate-dependent glutaminase activity in guinea pig and chicken, species with no net uptake of glutamine from blood, is also consistent with the role proposed for the enzyme. The correlation between blood glutamine concentration and glutamine uptake (Table I) suggests a possible adaptation of blood levels of glutamine related to the metabolic activity of the intestine or adaptation of intestinal metabolism to blood levels of glutamine. It may be noteworthy that the net uptake of glutamine by rat intestine is proportional to its concentration in plasma over the range 0.20.6 mM (1). The low activity observed or reported for several intestinal amidotransferases suggests that these enzymes, although they may be qualitatively important for the metabolism and development of intestinal epithelial cells, do not contribute significantly to the observed metabolism of glutamine. The contribution of “phosphateindependent, maleate-stimulated glutaminase” (46) depends upon how this activity is defined. Both phosphate-dependent glutaminase and “phosphate-independent, maleate-stimulated glutaminase” are believed to play an important role in glutamine hydrolysis in the kidney. In rat kidney, the latter activity is associated with y -glutamyltranspeptidase (7, 8)) which in the presence of maleate, has 5-10% as much phosphate-independent glutaminase as transpeptidase activity. Intestinal mucosa also contains y-glutamyltranspeptidase (1 pmol/h/mg of protein) (47). The activity is low relative to kidney and is enriched in the brush border of villus cells (48), where activity approaches 6 cl.mol/h/

GLUTAMINASE

IN

INTESTINAL

mg of protein (49). p-Hydroxymercuribenmate does not inactivate kidney “phosphate-independent, maleate-stimulated glutaminase” (25) or transpeptidase activity (50). Similarly, intestinal transpeptidase activity is not inactivated by p-hydroxymercuribenzoate (0.5 mM) or by chloroketone (20 mM) (L. Pinkus, unpublished observation). We found that in glutaminase assays of rat mucosal homogenates or mitochondria, the replacement of inorganic phosphate in the incubation mixture with 67 mM maleate led to 60-70% as much activity as with phosphate (Table III), but the addition of maleate to saturating concentrations of phosphate did not further stimulate activity. Furthermore, preincubation with either 0.5 mM p-hydroxymercuribenzoate or 20 mM chloroketone resulted in loss of more than 98% of the maleate-stimulated as well as the phosphate-stimulated activities. Thus, on the basis of its high activity, its mitochondrial localization, and its susceptibility to inactivation byp-hydroxymercuribenzoate and chloroketone, nearly all of the “phosphateindependent, maleate-stimulated glutaminase” activity in intestine is not due to y-glutamyltranspeptidase but is apparently due to phosphate-dependent glutaminase. Additional evidence that any transpeptidase-associated glutaminase activity of the brush border does not play a major role in hydrolyzing intestinal glutamine comes from the experiment in which chloroketone was administered luminally (Table V). Although this enzyme is not inhibited by chloroketone, glutamine administered subsequent to the inhibitor reached the venous blood without detectable hydrolysis. Katunuma et al. (25), without providing experimental details, have also reported the absence of “phosphateindependent glutaminase” from rat intestine. Glutamine hydrolysis in the epithelial cell mitochondria generates glutamate, which serves as a respiratory substrate, while most of the ammonia generated is released into portal venous blood. The rapid elimination of the ammonia maintains the tissue ammonia concentration, 0.74 pmol/g, at a level similar to the 0.59

MUCOSA

515

pmol/g observed in liver (l), a tissue that efficiently fixes ammonia. The intestinal glutaminase is ideally positioned to provide ammonia subsequently utilized by the liver. The rather high apparent Km reported for liver glutaminase, 28 mM (51), creates some uncertainty about the activity of this enzyme in uiuo. Furthermore, there appears to be little, if any, net uptake of circulating glutamine by the liver (1, 52). Thus, ammonia produced by intestinal glutaminase may be a more important nitrogen source for liver than ammonia generated by hepatic glutaminase. As in intestine, phosphate-dependent glutaminase is a mitochondrial enzyme in kidney (38, 53), liver (54), and Ehrlich ascites cells (55). The crude intestinal enzyme resembles the glutaminases of kidney, brain, and ascites cells in several respects. The apparent Km for glutamine, 2.2 mM, is similar to that reported for the phosphate-dependent glutaminase of kidney, 5 mM (40, 56,57), brain, 1.8-8 mM (58, 59), and Ehrlich ascites cells, 4.5 mM (55), and significantly different from that reported for the liver enzyme, 28 mM (51). The K,, for phosphate, 22 mM, is similar to the 25 mM value noted for the Tris form of the kidney enzyme (59), and the pH activity profile determined in phosphate buffer is like that reported for the “phosphate-borate” form of the kidney (40, 59) and brain enzymes (60). Activation by phosphate-containing compounds as well as inorganic and organic anions has also been noted for the enzymes of kidney and brain (40, 58, 61). Immunoprecipitation studies using antibody to the purified kidney glutaminase indicate that the enzymes from kidney and brain are identical immunologically and different from the liver enzyme (62). The glutaminases solubilized from kidney and intestinal mitochondria give identical immunoprecipitation curves with this antibody (N. Curthoys, personal communication). One significant difference between rat kidney and intestinal enzymes is their adaptive response during acidosis. Whereas the kidney enzyme increases in both its activity and quantity (62) during acidosis, the intestinal glutaminase activ-

PINKUS

516

AND WINDMUELLER

ity, like that of brain and liver (63), does not increase. During the course of our studies, Anderson et al. (64) reported that intestine from normal and acidotic rats produced ammonia at similar rates when incubated with glutamine in vitro. This is consistent with our observations on glutaminase activity. ACKNOWLEDGMENTS We thank Mr. Albert Spaeth for his technical assistance in the perfusion experiments and Mr. Joseph Pearce, Dr. Frank Macri, and Dr. James Vickers for providing some of the larger animals. Thanks are also given to Drs. Paul Richman, Ron Sekura, and M. Earl Balis for critical reading of the manuscript. 1. WINDMUELLER,

H. G., AND SPAETH, A. E. (1974). J. Biol. Chem. 249, 5070-5079. 2. WINDMUELLER, H. G., AND SPAETH, A. E. (1975) Arch. Biochem. Biophys. 171, 662-672. 3. FELIG, P., WAHREN, J., KARL, I., CERASI, E., LUFT, R., AND KIPNIS, D. M. (1973) Diabetes

5.

6. 7. 8. 9.

22, 573-576. PINKUS, L., AND MEISTER, A. (1972) J. Biol. Chem. 247, 6119-6127. STERN, B. K. (1966) Gastroenterology 51, 855866. WEISER, M. (1973) J. Bill. Chem. 248,2536-2541. TATE, S. S., AND MEISTER, A. (1975) J. Bio.!. Chem. 250, 4619-4627. CURTHOYS, N. P., AND KUHLENSCHMIDT, T. (1975) J. Biol. Chem. 250, 2099-2105. ORLOWSKI, M., AND MEISTER, A. (1965) J. Biol. Chem.

240, 338-347.

10. TATE,

S. S., AND MEISTER,

Chem. 11. SMITH,

249, 7593-7602. L. (1955) Methods

Chem.

27. 28. 29.

tarnine Metabolism (Prusiner, S., and Stadtman, E. R., eds.) p. 227-258, Academic Press, New York. KUPCHIK, H. Z., AND KNOX, W. E. (1970) Arch. B&hem. Biophys. 136, 178-186. TATIBANA, M., AND SHIGESADA, K. (1972) Adv. Enzyme Regul. 10, 249-271. PRAJDA, N., MORRIS, H. P., AND WEBER, G. (1976) Cancer Res. 36, 4639-4646. KAUFMAN, M., YIP, C., AND KNOX, W. E. (1971) Enzyme

12, 537-544.

30. KIKUCHI, H., KOBAYASHI, Y., AND TSUIKI, S. (1971) Biochim. Biophys. Actu 237,412-421. 31. POGELL, B. M., AND GRYDER, R. M. (1957) J. Biol. Chem. 228, 701-712. 32. WRISTON, J. C., JR. (1971) in The Enzymes (Boyer, P. D., ed.), 3rd ed., Vol. 4, p. 101, Academic Press, New York. 33. LEBLOND, C. P., AND STEVENS, C. E. (1948)Anat. Rec. 100, 357-377. 34. NORDSTROM, C., DAHLQUIST, A., AND JOSEFSSON, L. (1968) J. Histochem. Cytochem. 15,713-721. 35. IMONDI, A. R., BALIS, M. E., AND LIPKIN, M.

A. (1974) J. Biol.

(1969)

Exp.

Cell

Res.

58, 323-330.

36. FORSTNER, Biochem.Anul.

2,427-

37.

434. 12. WILLIAMS,

troenterology 40, 423-426. 24. HOROWITZ, M. L., AND KNOX, W. E. (1968) Enzymol. Biol. Clin. 9, 241-255. 25. KATUNUMA, N., KATSUNUMA, T., TOWATARI, T., AND TOMINO, I. (1973) in The Enzymes of Glu-

26.

REFERENCES

4.

Methods of Enzymatic Analysis (Bergmeyer, H. U., ed.), 2nd revised printing, pp, 384-388, Academic Press, New York. 21. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951)J. Biol. Chem. 193, 265-275. 22. RICHTERICH, R., GOLDSTEIN, L., AND COHEN, E. (1958) Hoppe-Seyler’s 2. Physiol. Chem. 312, 45-50. 23. SPENCER, R. P., AND ZAMCHECK, N. (1961) Gas-

C. H., AND KAMIN,

H. (1962) J. Biol.

237, 587-595.

13. COOPER, A. J. L., AND MEISTER, A. (1972) Biochemistry 11, 661-671. 14. TATIBANA, M., AND~HIGESADA, K. (1972) J. Biothem. (Tokyo) 72, 537-547. 15. KATUNUMA, N., AND WEBER, C. (1974) FEBS Lett. 49, 53-56. 16. LONG, C. W., LEVITSKI, A., AND KOSHLAND, D. E. (1970) J. Biol. Chem. 245, 80-87. 17. WINTERBURN, P. J., AND PHELPS, V. G. (1971) B&hem. J. 121, 701-709. 18. KLEMPERER, H. G., AND HAYNES, G. R. (1968) Biochem. J. 108, 541-546. 19. BREITMAN, T. R. (1963) Biochim. Bzbphys. A&

67, 153-158. 20. BERNT, E., AND BERGMEYER,

H.

U.

(1965)

in

38. 39. 40. 41. 42.

G. G., SABESIN, S. M., AND ISSELBACHER, K. J. (1968) B&hem. J. 106,381-390. WANG, T. Y. (1967) in Methods in Enzymology (Grossman, L., and Moldave, K., eds.), Vol. 21, pp. 417-421, Academic Press, New York. CURTHOYS, N. P., AND WEISS, R. F. (1974) J. Biol. Chem. 249, 3261-3266. GOLDSTEIN, L. (1966)Amer. J. Physiol. 210,661666. CURTHOYS, N. P., KUHLENSCHMIDT, T., AND GODFREY, S. S. (1976) Arch. Biochem. Biophys. 174, 82-89. WINDMUELLER, H. G., AND SPAETH, A. E. (1976) Arch. Biochem. Biophys. 175, 670-676. PINKUS, L. (1977) in Methods in Enzymology

(Jakoby, W. B., and Wilchek, M., eds.), Vol. 46, pp. 414-427. Academic Press, New York. 43. BUCHANAN, J. M. (1973) Adv. Enzymol. 39, 91183. 44. DAVIES, B. M. A., AND YUDKIN, J. (1952) Bio-

GLUTAMINASE

IN

them. J. ‘52, 407-412. 45. CURTHOYS, N. P., AND LOWRY, 0. H. (1973) J. Biol. Chem. 248, 162-168. 46. KATUNUMA, N., TOMINO, I., AND NISHIRO, H. (1966) Biochem. Biophys. Res. Commun. 22, 321-328. 47. GARVEY, T. Q., HYMAN, P. E., AND ISSELBACHER, K. J. (1976) Gastroenterology 71, 778-785. 48. CURTHOYS, N. P., AND SHAPIRO, R. (1975) FEBS Lett. 58, 230-233. 49. CORNELL, J. S., AND MEISTER, A. (1976) Proc. Nat. Acad. Sci. USA 73, 420-422. 50. SZEWCZUK, A., AND CONNELL, G. E. (1965) Biochim. Biophys. Acta 105, 352-367. 51. HUANG, Y. Z., AND KNOX, W. E. (1976) Enzyme 21, 408-426. 52. AIKAWA, T., MATSUTAKA, H., YAMAMOTO, H., OKUDA, T., ISHIKAWA, E., KAWANO, T., AND MATSUMURA, E. (1973) J. Biachem. (Tokyo) 74, 1003-1017. 53. KALRA, J., AND BROSNAN, J. T. (1974) J. Biol. Chem. 249, 3255-3260. 54. KALRA, J., AND BROSNAN, J. T. (1973) FEBS

INTESTINAL

MUCOSA

517

Lett. 37, 325-328. 55. KOVACEVIC, Z. (1974)CancerRes. 34,3403-3407. 56. KLINGMAN, J. D., AND HANDLER, P. (1958) J. Biol. Chem. 232, 369-380. 57. SAYRE, F. W., AND ROBERTS, E. (1958) J. Bial. Chem. 233, 1128-1134. 58. SVENNEBY, G. (1972) J. Neurochem. 19,165-174. 59. SVENNEBY, G., TVEIT, B., AND KVAMME, E. (1970) J. Biol. Chem. 245, 1878-1882. 60. SVENNEBY, G. (1970) J. Neurochem. 17, 15911599. 61. WEIL-MALHERBE, H. (1972) J. Neurochem. 19, 2257-2267. 62. CURTHOYS, N. P., KUHLENSCHMIDT, T., GODFREY, S. S., AND WEISS, R. F. (1976) Arch. Biochem. Biophys. 172, 162-167. 63. PITIX, R. F. (1973) in The Handbook of Physiology (Orloff, J., and Berliner R. W., eds.) Vol. 8, pp. 455-496, American Physiological Society, Washington, D. C. 64. ANDERSON, N. M., BENNETT, F. I., AND ALLEYNE, A. 0. (1976) Biochim. Biophys. Acta 437, 238-243.