Regulation of bacterial nitrogen assimilation by glutamine synthetase

TIBS - January

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1977

placement currents of the type depicted in Fig. IA, that is to say with the linear component of the capacity current independent of the membrane potential, we have seen that Qon remains equal to Qoff regardless of the duration of the pulse up to 32 ms at 6” C. However, currents of the type depicted in Fig. 1B are frequently recorded during the first few minutes after the onset of the perfusion. Studying the changes in displacement currents as a function of time we have seen that during the first 10 min, current transients for hyperpolarising voltage-clamp pulses (from a holding potential near - 100 mV) could be analysed in terms of two exponentially declining components. The first component Icl is terminated in less than 10 ,us at 6” C. The second, ICz, decays with a relaxation time constant smaller than 100 ps. With time (less than 60 min) a third exponential component became apparent, I,,. It had time constants ranging between 300 and 600 ps at 6”C. The charge transferred through each of these three components of the membrane capacity could be evaluated by measuring the records taken after 10 min and after 60 min under intracellular perfusion at 6” C. The charge transferred during Zc, (representing a membrane capacity of nearly 0.9 pF/cmZ) did not change with time. The charge transferred during the second component Zc, increased as a sigmoidal function of membrane potential and amounted to 0.45 pF/cmZ at midpoint. The charge transferred during the third component, Zcj, changed with membrane potential; for - 150 mV it amounted to nearly 0.5 pF/cm2. It is this third component of the displacement current that changes most with time under intracellular perfusion. Our interpretation is that Zc3 results from changes in the proteins at the inner edge of the axolemma. If protein groups are exposed and become free to ionize, the dielectric properties of the internal membrane interface will be changed drastically. The question of whether the on response obeys first or higher order kinetics has been studied systematically only in the case of myelinated nerve fibres [ 111.The results presented, examples of the several hundred records submitted to least-square fittings procedures, showed that the net charge transfer during and after pulses of equal size can be analysed in terms of a single exponential component. It should be pointed out here that with the data-acquisition system used in these experiments the maximum error in the determination of the charge is about 8 % of the average maximum charge displaced in the myelinated nerve tibre. Therefore, the maximum con-

tribution of any other non-exponential component to the charge Q(t) will be 8 % of the maximum charge. This conclusion supports the notion that the charge movement is of first order transition and bears important implications as to the mechanism. Organization of sodium channels

We propose that sodium channels are not permanent structures but functional aggregates of integral proteins (proteins completely or partially embedded in the lipid matrix of the membrane, and held in the membrane mainly by hydrophobic interactions). We postulate that channel formation involves the following components: (1) Selectivity filter or amphipathic proteins (integral proteins wjth two parts, one hydrophilic which protrudes from the membrane, and the other hydrophobic, which is embedded in the lipid matrix [2]) located on the outside of the membrane which serve as organisers of the sodium channel (see Fig. 3A). We identify this protein with the tetrodotoxin-binding protein of Levinson and Ellory with a molecular weight of 200,000. (2) Gating molecules or hydrophobic proteins (Fig. 3B) with a large dipole moment. From the relaxation time constant data for gating currents we estimate the molecular weight of 100,000 for these subunits.

Once the sodium channel is formed (Fig. 3C) the size of the aggregate is 500,000. The energy barriers within the pore formed between the segment of the organizer protruding into the hydrophobic part of the membrane and the gating units will determine whether Na+ could go; through the driving force to set this motion being the electrochemical potential gradent for Na+ (Fig. 3D). References 1 Cole, K. S. (1968) in Membranes,

Ions and Impulses - A Chapter of Classical Biophysics, University

of California Press, Berkeley 2 Zwaal, R. F. A., Demel, R. A., Roelofsen, B. and Van Deenen, L. L. M. (1976) Trends Biochem. Sci. 1, 112-114 3 Hodgkin, A. L. and Huxley, A. F. (1952) J. Physiol.

117,500-544

4 Armstrong, C.M., Bezanilla, F. and Rojas, E. (1973) J. Gen. Physiol. 62,375-391 5 Hille, B. (1970) Progr. Biophys. Mol. Biol. 21, 1-32 6 Bezanilla, F., Rojas, E. and Taylor, R. E. (1970) J. Physiol. 207, 151-164

Verveen, A. A. and DeFelice, L. J. (1974) Progr. Biophys. Mol. Biol. 28, 189-265 Keynes, R. D., Ritchie, J.M. and Rojas, E. (1971) J. Physiol. 213,253-254

Keynes, R.D., Bezanilla, F., Rojas, E. and Taylor, R.E. (1975) Phil. Trans. R. Sot. Land. Ser. B 270, 365-375 10 Nonner, W., Rojas, E. and Stimpfli, R. (1975) Pfltigers Arch. 354, 1-18

11 Rojas, E. (1976) Cold Spring Harbour Quant. Biol. XL, 305-320

Symp.

Regulation of bacterial nitrogen assimilation by glutamine synthetase Boris Magasanik The synthesis of enzymes capable ofproviding enteric bacteria with ammonia andglutamate is activated by glutamine synthetase. Glutamine synthetase regulates its own synthesis in response to the intracellular levels of2-ketoglutarate andglutamine which in turn depend on the extracellular concentration of ammonia and the metabolic state of the cell.

The preferred nitrogen source for enteric bacteria (Escherichia coli, Klebsiella aerogens, Salmonella typhimurium, etc.) is ammonia. Substitution of another source of nitrogen for ammonia in a minimal medium with glucose as the source of carbon almost invariably results in slower growth. The principal product of ammonia assimilation is glutamate. This compound in turn is the precursor of several amino acids and furnishes the B.M. is Professor of Microbiology at the Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, U.S.A.

amino group of others by transamination. It is also the precursor of glutamine whose amide group provides some of the nitrogen atoms of amino acids and or purine and pyrimidine nucleotides. Other nitrogencontaining compounds can substitute for ammonia in the growth medium to support the growth of cells that are equipped with enzymes capable of converting them to ammonia or glutamate. It is immediately apparent that such enzymes fulfill a useful function in cells deprived of ammonia. It is therefore not surprising to discover that their rapid syn-

TIBS - January 1977

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thesis occurs only in the absence of ammonia. Recent studies have shown this regulation of the synthesis of these enzymes to be mediated by glutamine synthetase.

ADENY LY LATION

GS

PIIA

Active

*GS-AMP Inactive

Glutamine synthetase and ammonia assimilation Glutamine synthetase plays an essential role in ammonia assimilation for cells growing in media whose concentration of NH: is below 1 mM. The enzyme long considered responsible for the conversion of ammonia to glutamate, glutamate dehydrogenase (EC 1.4.1.4), fails to fulfill this role at low ammonia concentrations because the reaction it catalyzes has an unfavorable equilibrium for glutamate formation [1,2]. NH,+ 2-ketoglutarate + NADPH + H +--) *glutamate + NADP + + H,O

(1)

When the extracellular concentration of ammonia is low, glutamate is formed by the combined action of glutamine synthetase and glutamate synthase [ 1,2]. Glutamate+ ATP+ NH,+ --t glutamine + ADP + Pi

(2)

Glutamine + 2-ketoglutarate + NADPH + H ++ +2-glutamate+NADP+

(3)

Regulation of glutamine synthetase activity Cells growing in a medium with ammonia in low concentration contain active glutamine synthetase; addition of ammonia to the growth medium results in rapid inactivation of the enzyme’s ability to form glutamine; the inactive enzyme retains glutamyltransferase activity and can thus be measured readily. Removal of excess ammonia from the growth medium results in the rapid conversion of the inactive enzyme to the active form [3,4]. The mechanism of activation and inactivation of glutamine synthetase of E. coli has been elucidated by Stadtman and his collaborators [5] (Fig. 1). Glutamine synthetase is a dodecamer composed of identical subunits and has a molecular weight of approx. 600,000. The enzyme is inactivated by the attachment of adenylyl groups to a maximum of 12, one to a tyrosine residue on each of the subunits. The adenylylation requires ATP and two proteins called adenylyltransferase (ATase) and Pn. The same two proteins arenalso required for the removal of the adenylyl groups, which restores the biosynthetic function of glutamine synthetase. Whether adenylyltransferase and Pn catalyze adenylylation or deadenylylation depends on the form of PII. The attachment of uridylyl groups to this protein converts it

PIID GS (gin A)

- UMP

GS

- AMP

A6P

form. Conversely, when the ammonia concentration is high, the level of 2-ketoglutarate is reduced by its conversion to glutamate, and the level of glutamine increases. Under this condition, the conversion of PII* to PIID is not activated, and presumably PnD loses its uridylyl groups and is converted to PEA; in addition, adenylylation by adenylyltransferase . PIIA is stimulated: the result is conversion of glutamine synthetase to the inactive adenylylated form. It is of interest that glutamine synthetase plays a role in its own regulation: it catalyzes the synthesis of glutamine, which, as we have seen, stimulates its conversion to the inactive form.

DEADENYLYLATION

Fig. I. Adenylylation and deadenylylation of glutamine synthetase, adaptedfrom Ginsburg and Stadtman [5]. GS, glutamine synthetase (EC 6.3.1.2) ; ATase, adenylylation enzyme; UTase, uridylyl-removing enzyme; gln=gene controlling production of glutamine synthetase.

from PnA which activates adenylylation, to Pno which activates deadenylylation. The uridylylation of Pn is catalyzed by an enzyme called uridylyltransferase (UTase). Another enzyme catalyzes the removal of uridylyl groups from PIID. Studies using cell extracts have shown that the activity of uridylyltransferase is regulated by 2-ketoglutarate and glutamine: the former activates and the latter inhibits this enzyme. Conversely, glutamine activates and 2-ketoglutarate inhibits adenylylation by the complex adenylyltransferase’PIIA. We can see therefore that in the intact cell growing with glucose as source of carbon, adenylylation and deadenylylation of glutamine synthetase are controlled by the concentration of ammonia in the growth medium. When the ammonia concentration is low, 2-ketoglutarate will accumulate in the cells and glutamine will be in short supply. Under this condition, adenylylation by adenylyltransferase . PIIA is inhibited; conversion of PEA to PIID by uridylyltransferase is stimulated; the result is conversion of glutamine synthetase to the active non-adenylylated

Regulatiou of glutamine synthetase synthesis The composition of the growth medium determines not only the activity but also the intracellular level of glutamine synthetase. The highest levels of the enzyme are formed in cells of E. co/i or K. aerogenes growing in media containing glucose as major source of carbon, and either ammonia in growth-rate limiting concentration, or a poor source of nitrogen, such as an amino acid or nitrate in place of ammonia [6]. In cells growing in media containing glucose and an excess of ammonia, the levels of glutamine synthetase are six-ten-fold lower. A further five-ten-fold reduction of the enzyme level can be obtained in K. aerogenes by substituting histidine, a poor source of energy, for glucose and adding glutamine to the ammonia-containing medium (Bender, R. A. and Magasanik, B., unpublished observation). Thus the level of glutamine synthetase can vary 50-loo-fold in response to changes in the environment of the cell. This variation in enzyme level is the result of regulation of transcription of the structural gene (glnA) for glutamine synthetase; the enzyme level reflects the intracellular concentration of messenger RNA complementary to glnA-specific DNA (Weglenski, P. and Tyler, B.M., unpublished

TABLE I Regulation of synthesis of enzymes of nitrogen metabolism in Klebsiella Enzyme(s)

Substrate

Products

Regulation by Inducer CAP*-cyclic AMP

hut Put nut dut

Nitrogenase Urease Asparaginase Tryptophan TA

Histidine Proline Arginine Putrescine N, Urea Asparagine Tryptophan

Glutamate, NH, Glutamate Glutamate, NH, Glutamate NH, CO,, NH, Aspartate, NH, Glutamate

+ + + + -

+ + + + _ _ -

*Cyclic AMP receptor protein or catabolite-gene activator protein.

Glutamine synthetase + + + + + + + +

11

TIBS - January 1977 observation). Glutamine synthetase itself plays a key role in the regulation of the transcription of its structural gene. This conclusion stems from the investigation of glutamine-requiring mutants of K. aerogenes. These mutants contain no or very little glutamine synthetase. The mutations occur at three unlinked sites on the chromosome gInA, glnB, and glnD. Mutations ingZnA and inglnE, a site linked to glnB, restore to a mutant in glnB the ability to produce glutamine synthetase [6,7,81. The demonstration that some glnA mutants produce enzymatically inactive glutamine synthetase antigen or heatlabile enzyme identilies glnA as site of the structural gene for glutamine synthetase [9]. Examination of extracts of mutants in glnB, glnD and glnE revealed each to be defective in a different component of the adenylylation/deadenylylation system. The glnD mutation resulted in a loss of uridylyltransferase activity and the glnB mutation has altered PI* so that it cannot assume the PIID form required for stirnulation of deadenylylation. These mutations leading to the inability of the cell to deadenylylate glutamine synthetase result in abnormally low levels of the enzyme. The mutation in glnE has altered the adenylyltransferase so that it fails to catalyze the adenylylation of glutamine synthetase, but catalyzes its deadenylylation whether combined with PnD or PuA. This failure to adenylylate glutamine synthetase results in high levels of unadenylylated enzyme even in cells grown with an excess of ammonia (GlnC - phenotype). Together these results suggest the possibility that the adenylylated form of glutamine synthetase represses glutamine synthetase. This hypothesis receives support from the observation that mutations in glnA, the structural gene for glutamine synthetase, suppress the mutation in glnB and also result in the GlnC- phenotype [71. The introduction of an episomal glnA + gene into cells carrying the mutated glnA gene responsible for GlnC- phenotype, restores almost normal regulation of glutamine synthetase synthesis [lo]. It would appear that the mutation in the chromoso-G ma1 glnA gene has altered glutamine synthetase so that it fails to repress glutamine synthetase while retaining enzymatic activity. The normal glutamine synthetase determined by the episomal glnA + gene restores repression. Although the hypothesis that adenylylated glutamine synthetase is the repressor accounts satisfactorily for the results obtained so far, it cannot be ruled out that non-adenylylated glutamine synthetase is an activator of the formation of glutamine synthetase. In that

case the apparent dominance of glnA+ responsible for the GlnC+ phenotype could be due to subunit mixing. It is also possible that the synthesis of glutamine synthetase is activated by unadenylylated and repressed by adenylylated glutamine synthetase. Regulation by glutamine synthetase

The fact that both amount and form of glutamine synthetase change in response to the ammonia concentration of the medium offers a sensible explanation for the role of glutamine synthetase in the regulation of the synthesis of enzymes capable of providing the cell with glutamate or ammonia in K. aerogenes. The enzymes of the hut system catalyze in four steps the conversion of L-histidine to glutamate, ammonia and formamide. The formation of these enzymes requires the presence of histidine and the absence of either glucose or ammonia. The presence of histidine is necessary for the inactivation of a repressor capable of binding to hut-specific DNA and preventing its transcription. The absence of the good energy source, glucose, permits an increase in the intracellular level of 3’:.5’-cyclic AMP which combines with a protein (CAP) and stimulates the transcription of the hut genes [l l-131. The absence of ammonia leads to an increase in unadenylylated glutamine synthetase, which, like CAP charged with cyclic AMP, stimulates transcription of hut [14]. In this manner, the formation of the system is regulated according to the metabolic needs of the cell. The hlct enzymes are only produced when histidine is available and when the cell either requires energy or nitrogen. The existence of mutants unable to produce glutamine synthetase (GlnA-) or producing glutamine synthetase at a high level even in the presence of ammonia (GlnC -) has made it possible to determine which systems are regulated by glutamine synthetase. GlnC- mutants produce the enzyme of the hut system even in the presence of glucose and ammonia; GlnAmutants fail to produce these enzymes even during ammonia starvation [7]. An exception are mutants in gInA, producing enzymatically inactive glutamine synthetase antigen at a high level even in the presence of ammonia. These glutamine-requiring mutants produce the enzymes of the hut system in the presence of glucose and ammonia. It appears that the enzymatically inactive glutamine synthetase has retained the ability to stimulate transcription of hut [7]. The results obtained with other enzymes are presented in Table I. Enzymes capable of supplying the cell with energy and with glutamate, for example those degrading

proline [13], arginine or putrescine (Friedrich, B. and Magasanik, B., unpublished observation), are regulated like hut by specific induction, cyclic AMP and glutamine synthetase. Other enzymes capable only of supplying the cell with glutamate or ammonia such as urease [ 151, nitrogenase [ 161, tryptophan transaminase (Paris, G. and Magasanik, B., unpublished observation) and asparaginase [ 171do not require induction, do not respond to cyclic AMP and are exclusively regulated by glutamine synthetase. In all those cases glutamine synthetase stimulates production of the enzymes. In addition, glutamine synthetase inhibits the formation of glutamate dehydrogenase [18]. This enzyme plays no role in ammonia-starved cells. Operation of the system

Let us consider cells growing in a medium containing glucose, histidine and ammonia. Initially, these cells use glutamate dehydrogenase to assimilate ammonia; they contain little glutamine synthetase, and that mostly in an adenylylated form and consequently unable to activate transcription of the genes coding for the histidine-degrading enzymes (hut). We assume that these cells have used up most of the ammonia, so that its level has fallen below 1 mM. Now, glutamate dehydrogenase will no longer be able to function and the cellular level of 2-ketoglutarate will increase; glutamine synthetase, present largely in the inactive form will not function effectively, and the cellular level of glutamine will decrease. The consequent increase in the ratio of 2-ketoglutarate to glutamine will cause deadenylylation of glutamine synthetase, and thus remove the repressor of glutamine synthetase. As a glutamine result, non-adenylylated synthetase will accumulate in the cell, repress glutamate dehydrogenase and activate the transcription of the hut system. The degradation of histidine by the hut enzymes will provide glutamate as well as ammonia, which in turn will be converted to glutamate through combined action of and glutamate glutamine synthetase synthase. Let us now supply these cells, after several generations of growth on glucose and histidine, with an excess of ammonia. Because these cells are rich in active glutamine synthetase, this will result in an increase in the intracellular level of glutamine. The consequent decrease in the ratio of 2-ketoglutarate to glutamine will lead to adenylylation of glutamine synthetase, removing the activator of hut transcription and the repressor of glutamate dehydrogenase, and providing the repressor of glu-

TIBS - January 1977

12

tamine synthetase. As a result, the cells will cease to produce the hut enzymes and glutamine synthetase, and begin to produce glutamate dehydrogenase. After several generations of growth in this medium, they will have acquired the enzymatic constitution appropriate for growth in excess ammonia. Acknowledgements The authors are recipients of Public Health Service research grants GM-07446 from the National Institute of General Medical Sciences and AM- 13894 from the National Institute of Arthritis and Metabolic Diseases, and grant GB-5322 from the National Science Foundation.

5 Ginsburg, A. and Stadtman, E. R. (1973) in The Enzymes of Glutamine Metabolism (Prusiner, S.

and Stadtman, E. R., eds), pp. 9-43, Academic Press, New York 6 Prival, M. J., Brenchley, J. E. and Magasanik, B. (1973) J. Biol. Chem. 248,4334-4344 7 Streicher, S. L., B:nder, R. A. and Magasanik, B. (1975) J. Bacferiol. 121, 32&331 8 Foor, F., Janssen, K. A. and Magasanik, B. (1975) Proc. Nat. Acad. Sci. U.S.A. 72, 4844.4848 9 DeLeo, A.B. and Magasanik, B. (1975) J. Bacteriol. 121,313-319

10 Streicher, S. L., DeLeo, A.B. and Magasanik, B. (1976) J. Bacterial. (in press) 11 Smith, G.R. and Magasanik, B. (1971) J. BioL Chem. 246,3330-3341

12 Hagen, D. C. and Magasanik, B. (1973) Proc. Nat. Acad. Sci. U.S.A. 70, SOS-812

13 Prival, M.J. and Magasanik, B. (1971) J. Biol. Chem. 246,62886296

14 Tyler, B., DeLeo, A. B. and Magasanik, B. (1974) Proc. Nat. Acad. Sci. U.S.A.

References

71,2_25-229

15 Friedrich, B. and Magasanik, B. (1976) Abstr.

1 Meers, J.L., Tempest, D. W. and Brown, C.M. (1970) J. Gen. Microbial. 64, 187-194 2 Tempest, D. W., Meers, J.L. and Brown, C.M. (1970) Biochem. J. 117,405-407 3 Wulff, K., Mecke, D. and Holzer, M. (1967) Biothem. Biophys. Res. Commun. 28, 740-745 4 Kingdon, M.S., Shapiro, B.M. and Stadtman, E.R. (1967) Proc. Nat. Acad. Sci. U.S.A. 58, 1703-1710

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16 Streicher, S. L., Shanmugam, K.T., Ausubel, F., Morandi, C. and Goldberg, R.B. (1974) J. Bacferiol. 120, 815-821 17 Resnick, A. D. and Magasanik, B. (1976) J. Biol. Chem. 251,2722-2728

18 Brenchley, J. E., Prival, M. J. and Magasanik, B. (1973) J. Biol. Chem. 248,6122-6128

depletion of lactose, they rise again. This wave-like fluctuation of the levels during repeated cycles of growth shows the tendency for cyclic AMP to be low during periods of growth and high during periods of stasis or growth transition. The physiological significance of these alterations appears to be that the adaptation to a new growth situation always requires higher cyclic AMP levels than does the continued growth on that carbon source. It is likely that raised levels of cyclic AMP overcome the barrier to effective transcription of operons for catabolic enzymes when only low levels of inducer penetrate the cells. After induction of the transport system for a catabolite, effective transcription can persist with lowered concentrations of cyclic AMP. The following discussion will show that the presence of a sugar-transport system provides the key to catabolite-dependent lowering of cyclic AMP levels. Interaction of a sugar-transport system with adenylate cyclase creates a complex that can regulate the enzyme activity. Sugars inhibit adenylate cyclase activity

Regulation of Escherichia coli adenylate cyclase by phosphorylationdephosphorylation Alan Peterkofsky The mechanism of catabolite repression involves a sugar-dependent regulation of adenylate cyclase via phosphorylation-dephosphorylation. This is accomplished by an interaction of adenylate cyclase with the sugar transport system.

In Escherichia coli, the regulation of the rate of synthesis of induced enzymes required for the degradation of most carbon sources calls for not only the presence of inducer but also an optimal concentration of cyclic AMP [1,2]. A major advance in our understanding of the control of induced enzyme synthesis was the notion that catabolites exerted a negative control over cellular cyclic AMP levels [I,31 (the phenomenon of catabolite repression). Recent studies have mapped out the essential characteristics of this process. It has been shown that sugars inhibit adenylate cyclase if the appropriate transport system for that sugar is induced [4]. A.P. is at the Laboratory of Biochemical Genetics, National Heart and Lung Institute, Bethesda, Maryland 20014, U.S.A.

The membrane-bound complex between adenylate cylcase and the sugar transport system permits the sugar-dependent regulation of adenylate cyclase by way of a phosphorylation-dephosphorylation mechanism [5]. When E. coli are presented with a growth medium containing a mixture of glucose and lactose, they grow sequentially on the glucose followed by the lactose (Fig. 1) (diauxic growth). An examination of the cellular cyclic AMP levels in such a situation showed that they rise abruptly during the transition period when cells are adapting from growth on glucose to growth on lactose. Just prior to the initiation of growth on lactose, the levels drop again and remain low throughout the period of lactose growth. When growth stops due to

While measurements of adenylate cyclase in broken cell preparations show no inhibition by glucose, intact or permeabilized cells are sensitive to glucose inhibition (Fig. 2) [6,7]. These observations have led to the idea that sugars do not inhibit adenylate cyclase directly but require some other factors, which have been suggested to be the sugar-transport systems [4,5]. When E. coli were grown on glucose, the inhibition of adenylate cyclase by glucose was relatively specific; most of the other stereoisomeric hexoses were inactive. Glucose 6-phosphate, which is the first product of glucose metabolism in E. coli, was also inactive, suggesting that the observed inhibition was due to free glucose itself. The absence of a requirement for glucose metabolism was also supported by the inhibitory effects of the glucose analogues 2-deoxyglucose and methyl-a-glucoside, neither of which support growth or are metabolized, although they are taken up by cells and phosphorylated. The first evidence that transport systems were involved in the regulation of adenylate cyclase activity came from the observation that the profile of sugars that inhibited adenylate cyclase varied with the growth conditions of the cells (Table I) [4]. Irrespective of the carbon source used for growth, adenylate cyclase was always inhibited by glucose whose transport system is partially constitutive. However, when cells were grown on a variety of other carbon sources, like fructose or mannitol, the adenylate cyclase from such cells was