The phosphorylated form of the enhancer-binding protein NTRC has an ATPase activity that is essential for activation of transcription

Cell,

Vol.

67, 155-167,

October

4, 1991,

Copyright

0 1991

by Cell

Press

The Phosphorylated Form of the Enhancer-Binding Protein NTRC Has an ATPase Activity That Is Essential for Activation of Transcription David S. Weiss, Jacques Batut,” Karl E. Klose, John Keener,t and Sydney Kustu Department of Plant Pathology and Department of Molecular and Cell Biology University of California Berkeley, California 94720

Summary The NTRC protein of enteric bacteria is an enhancerbinding protein that activates transcription in response to limitation of combined nitrogen. NTRC activates transcription by catalyzing formation of open complexes by RNA polymerase (9 holoenzyme form) in an ATP-dependent reaction. To catalyze open complex formation, NTRC must be phosphorylated. We show that phosphorylated NTRC has an ATPase activity, and we present biochemical and genetic evidence that NTRC must hydrolyze ATP to catalyze open complex formation. It is likely that all activators of 054 holoenzyme have an ATPase activity. Introduction In addition to the most abundant sigma factor, cr70,eubacteria employ alternative sigma factors that confer different promoter specificities on the core form of RNA polymerase. Whereas most alternative sigma factors allow transcription of genes whose products contribute to a common physiological response, ds4 differs in that it is needed for transcription of genes whose products have diverse physiological roles (for reviews see Kustu et al., 1989; Thony and Hennecke, 1989). c? is not homologous to other known sigma factors, and a54-dependent promoters are characterized by an unusual spacing of the recognition elements (-12, -24). o”-dependent genes appear to share a mechanism of transcriptional activation-transcription by os4 holoenzyme depends on an activator protein bound to sites located at a distance from the transcriptional start. The proteins that activate transcripiion by os4 holoenzyme have in common a domain of ~240 amino acid residues that appears to be directly responsible for transcriptional activation (Figure 1A). Moreover, all of the activators contain a recognizable nucleotide-binding motif within this domain (Ronson et al., 1987). The best-studied activator of o” holoenzyme is the nitrogen-regulatory protein NTRC of enteric bacteria (also called NRI). The mechanism of activation by NTRC has been studied intensively at the major promoter for the g/nA gene, which encodes glutamine synthetase. To activate

‘Present address: Laboratolre de Biologie Plantes-Microorganismes, CNRS-INRA, net-Tolosan, Cedex, France. fPresent address: Department of Biological California, Irvine, California 92717.

Moleculaire BP27, Chemistry,

des Relations F31326 CastaUniversity

of

transcription, NTRC binds to enhancer-like sites (Reitzer and Magasanik, 1986) located over 100 bases upstream of the start siteof transcription and contacts the polymerasepromoter complex directly by means of DNA loop formation (Reitzer et al., 1989; Su et al., 1990; Wedel et al., 1990). NTRC activates transcription by catalyzing the isomerization of closed recognition complexes between a54 holoenzyme and theg/nA promoterto open complexes, in which the DNA strands are locally denatured around the transcriptional start site (Sasse-Dwight and Gralla, 1988; Popham et al., 1989). The isomerization reaction requires ATP (Popham et al., 1989). Control of transcription at css4-dependent promoters appears to be accomplished primarily by modulating the function of the various activator proteins. For different activator proteins, function is modulated by different mechanisms and in response to different physiological signals (Kustu et al., 1989; Thony and Hennecke, 1989). The activity of NTRC is controlled positively by phosphorylation by NTRB (also called NRII) (Ninfa and Magasanik, 1986; Keener and Kustu, 1988; Weiss and Magasanik, 1988); phosphorylation increases under conditions of nitrogen limitation. The phosphorylation site in NTRC is within an amino-terminal domain of -120 residues (Figure 1A) (Keener and Kustu, 1988). A homolog of this domain has been found on more than 20 bacterial regulatory proteins whose activity is either known or thought to be modulated by phosphorylation (for reviews see Albright et al., 1989; Stock et al., 1989); these are the receiver proteins of “twocomponent” regulatory systems, which play a prominent role in signal transduction in eubacteria (Nixon et al., 1986; Kofoid and Parkinson, 1988). Little is known about the mechanism by which phosphorylation regulates the function of receiver proteins. Although phosphorylation of NTRC is not required for binding to its enhancer-like sites in the g/nA promoter regulatory region, it is absolutely required for its function as a transcriptional activator. We show that NTRC has an ATPase activity that is essential for open complex formation. We also show that a primary role for phosphorylation of NTRC is to stimulate its ATPase activity. We infer that other activators of 05” holoenzyme must also have an ATPase activity but that their ability to hydrolyze ATP will not be regulated by phosphorylation in every case. Results Phosphorylation of NTRC Activates an ATPase Activity We have previously demonstrated that ATP is required directly for the formation of open complexes by crw holoenzyme at the glnA promoter (Popham et al., 1989) as well as being required for phosphorylation of NTRC. To show a direct ATP requirement, we used a mutant form of NTRC (NTRCSIGOF[constitutive]) that has weak ability to activate transcription without being phosphorylated. (The activity of this mutant form, which has a lesion in its central domain

Cell 156

ATP

o0 Figure

1. Domain

structure

ADP

10

20 30 Time (mid

40

50

! 10

' 7 20 Time (mf$

40

1 so

of NTRC

(A) NTRC from S. typhimurium is 469 amino acids long and contains domains for regulation (-120 amino acids), transcriptional activahon (~240 amino acids), and DNA binding (-60 amino acids) (reviewed in Weiss et al., 1991). The regulatory domain is phosphorylated at aspartate 54 (circled “P”). The activation domain contains an ATPbinding motif (GESGSGK in one-letter amino acid code). The underlined glycine in the ATP-binding motif is at position 173 and is mutated to asparagine in one of the NTRCY’““‘“’ proteins (see below). The amino acids that are identical in all activators of os4 holoenzyme whose sequence is known are indicated in black. (6) Location of amino acid substitutions in an NTRCc”“s”‘““” protein (above) and several NTRC’“P’“““O’ proteins (below). Note that all repressor mutations occur in the transcriptional activation domain. With one exception they change residues that are identical in all of the activators whose sequence has been determined. (The exception: glutamate 208 is aspartate in FHLA.)

B

1

OJ 0 Figure

[Serl60-*Phe; Figure 1 B], is greatly increased by phosphorylation [Popham et al., 1989; Weglenski et al., 19891.) When studied in the absence of its cognate phosphotransferase NTRB, NTRCS’GOF still required ATP to catalyze open complex formation; ADP and analogs of ATP with nonhydrolyzable 8-r bonds failed to substitute (Popham et al., 1989). Since NTRC has a sequence motif found in many nucleotide-binding proteins, we hypothesized that NTRC might have to hydrolyze ATP to catalyze open complex formation. Consistent with this hypothesis, we have found that both wild-type NTRC and NTRCSIGOF(see below) have ATPase activity. For the wild-type protein this activity is dependent upon phosphorylation. We assayed the ATPase activity of NTRC-phosphate by quantitating release of inorganic phosphate, P,, under conditions in which NTRC-phosphate was generated in situ-in reaction mixtures containing NTRC, NTRB, and [yJ2P]ATP. We generated NTRC-phosphate in situ, rather than using purified material, because NTRC-phosphate has a rapid autophosphatase activity that makes it difficult to purify in high yield or maintain in constant amount during experiments (Figure 2A) (Keener and Kustu, 1988; Weiss and Magasanik, 1988). To detect ATPase activity during a phosphorylation reaction, it was necessary to distinguish P, released by the ATPase from P, released by the autophosphatase. In our first experiments we monitored P, release at early time points during a phosphorylation reaction-when the level of NTRC-phosphate was low-to minimize the contribution of the autophosphatase to the observed P, produc-

3 2

2. Time

Course

of Phosphorylation

of NTRC

(A) NTRC (1 hM) was phosphorylated by NTRB (100 nM) at an ATP concentration of 0.25 mM, as described in Experimental Procedures (see also Keener and Kustu, 1988). The time courseof phosphorylation (filled diamonds) was obtained by initiating a reaction with labeled MgATP (3360 cpmlpmol). After 20 min the autodephosphorylation rate was determined by adding a portion of the reaction mixture to prewarmed unlabeled MgATP to give a final ATP concentration of 13 mM (open circles). The half-life of NTRC-phosphate determined from these data is 3.2 min. The inset diagrams the phosphorylation and autophosphatase activities that underlie the time course of phosphorylation. (B) In the same experiment shown in (A), a second reaction (open diamonds) was initiated with unlabeled ATP at 0.25 mM and allowed to proceed for 10 min before addition of carrier-free [y-32P]ATP (final specific activity 3085 cpmlpmol). Samples were taken at the times indicated to monitor the rate at which the label equilibrated into NTRCphosphate.

tion. Control reactions showed that wild-type NTRC alone neither incorporated nor released much P, (Table 1A and see Table 3). Similarly, NTRB alone did not release much P, (Table 1A; NTRB is a phosphotransferase and incorporates P, under these conditions). In contrast, when NTRC and NTRB were present together, there was a high initial rate of P, incorporation, which is known to be largely into NTRC, and a higher rate of P, release (Table 1B). Since the rate of P, release exceeded the rate of incorporation, it could not be accounted for solely by the autophosphatase activity of NTRC-phosphate. (Recall that NTRC-phosphate is a$cumulating during the initial phase of the reaction [Figure 2A], so the rate of phosphorylation must exceed the rate of autodephosphorylation.) We hypothesized that the high initial rate of P, release was due, at least in part, to an ATPase activity of NTRC-phosphate.

Phosphorylated 157

Table

1. ATPase

Experiment

Proteins Conditions

A

NTRC NTRB

B

C

NTRC

Activity

Is an ATPase

of Phosphorylated

and

Rate of P, Incorporation (pmollminll0

NTRC

~1)

Rate of P, Release (pmoll min/lO ~1)


0.8” 0.2b

NTRC + NTRB Initiated with y32P-labeled ATP NTRC + NTRB Preincubated with unlabeled ATP

5.2

6.7”

2.5

40

NTRC + NTRB Preincubated N-terminus + NTRB Preincubated

1 .o

16

2.9

1.6

The initial rates of P, incorporation and P, release were determined as described in Experimental Procedures and are given in units of pmoll min/lO PI of reaction mixture. The endogenous 32P, in the [Y-~*P]ATP used in these experiments corresponded to a background of 30 pmol per IO ~1 of reaction mix. We could reliably detect P, release that was 10% over background. In experiment A, reactions contained 4 FM NTRC or 240 nM NTRB, which were warmed in buffer at 37V for 4 min before addition of unlabeled ATP to 0.25 mM. After 10 min incubation at 37%, carrierfree [yJ*P]ATP was added to a specific activity of 1506 cpmlpmol, and incubation was continued. P, release above background was first detected 30 min after addition of label and was linear for 120 min; the values reported are based on a 60 min time point. In experiment B, reactions contained 3.0 PM NTRC, 190 nM NTRB, 0.25 mM ATP (1600 cpmlpmol). Proteins were warmed in buffer (containing Mg2+ at 2 mM) at 37OC for 4 min. The first reaction was initiated by adding a prewarmed mix of labeled and unlabeled ATP. The second reaction was initiated by adding warmed unlabeled ATP. After 10 min of “preincubation” with unlabeled ATP to allow the level of NTRCphosphate to reach steady state (Figure 2B), warmed carrier-free [Y-~~P] ATP was added. Samples for measuring P, incorporation and P, release were taken 30 s after addition of label. In experiment C, reactions contained 140 nM NTRB and 2.3 PM NTRC, full-length or purified amino-terminal regulatory domain as indicated, and 0.25 mM ATP (1300 cpm/pmol). Manipulations were as for the “preincubation” tube of experiment B, except that with the amino terminus no P, release was detected after 30 s; the value reported was determined 9 min after addition of label. The rate determined for fulllength NTRC at 9 min was 14 pmol/min/lO ~1 of reaction mix. a The rate of ATP hydrolysis by unphosphorylated NTRC (experiment A) was (0.8 pmol/min/lO ~I)/(40 pmol NTRCllO ~1) = 0.02 pmollminl pmol NTRC. The rate of ATP hydrolysis by phosphorylated NTRC (experiment B) was (6.7 pmol/min/lO ~1)/(2.6 pmol NTRC-phosphate/ IO ~1) = 3 pmollminlpmol NTRC-phosphate. The 2.6 pmol NTRCphosphate is an average for the first minute of the experiment and assumes incorporation of only 1 mol of phosphate per dimer. b The observed incorporation is a minimum estimate, because the histidine-phosphate linkage in NTRB is labile to acid, which was used to precipitate phosphorylated proteins. The observed P, release is a maximum estimate, because it includes acontribution from acid hydrolysis of the histidine-phosphate linkage in NTRB.

To demonstrate the ATPase activity more convincingly, we increased the amount of NTRC-phosphate present in reaction mixtures. To do this we initiated phosphorylation reactions with unlabeled ATP and allowed 10 min for the level of NTRC-phosphate to reach steady state; steady state occurs when the rate of phosphorylation by NTRB is equalled by the rate of autodephosphorylation (Figure 2A). We then added [yJ2P]ATP and measured the initial rate of

P, release. Under these conditions (Table 1 B and see Table 3), the initial rate of P, release was dramatically higher than in the absence of preincubation with unlabeled ATP, although the rate of phosphorylation of NTRC was slightly lower. These results were consistent with the view that NTRC-phosphate has an ATPase activity. We employed this prephosphorylation protocol for studying the ATPase in most subsequent experiments. As a rough estimate of the degree to which phosphorylation stimulates the rate of ATP hydrolysis, we normalized the rate of P, release to the number of pmol of NTRC dimers (experiment A) or the number of pmol of P, incorporated into NTRC (experiments B and C). When adjusted in this fashion, the initial rate of ATP hydrolysis was about 0.02 pmol ATPlminlpmol of NTRC dimers in the absence of phosphorylation and about 3 pmol ATPlminlpmol of incorporated P, (Table 1). This corresponds to an activation of m200-fold. We have seen activation as great as several thousand fold (e.g., Table 3). Phosphorylated NTRC catalyzed release of ADP from [a-32P]ATP at the same rate as release of P, from [y-32. P]ATP (Figure 3A). This result showed that NTRCphosphate is indeed an ATPase rather than a phosphatase. The ATPase Activity Requires Full-Length NTRC We have previously shown that the kinetics of phosphorylation and autodephosphorylation of the isolated aminoterminal regulatory domain of NTRC (obtained following limited proteolysis with trypsin) are similar to those of the full-length protein (Keener and Kustu, 1988). We used the amino-terminal fragment to confirm directly that the high levels of P, release seen with full-length NTRC were not simply a consequence of the phosphorylation and subsequent dephosphorylation of NTRC. As expected, the rate of P, release by the phosphorylated regulatory domain was less than the rate of P, incorporation (Table 1 C). This result is congruent with our hypothesis that the ATPase activity of NTRC is associated with a nucleotide-binding motif in its central domain. The ATPase Activity Is Defective in Some Mutant NTRC Proteins That Fail to Activate Transcription Genetic evidence indicates that the ATPase activity of NTRC-phosphate is required for open complex formation: several mutant NTRC proteins that were isolated on the basis of their inability to activate transcription (Wei and Kustu, 1981) have little or no ATPase activity when phosphorylated (Table 2). These mutant proteins bind DNA normally and are phosphorylated and autodephosphorylated normally; they fail specifically in transcriptional activation (D. Weiss and S. Kustu, unpublished data). They are called NTRCrePreSsorproteins, because they retain the ability to repress transcription from a secondary o”‘-dependent promoter that lies in the enhancer region upstream of glnA (Reitzer and Magasanik, 1985). NTRC’eP’e”““’ proteins carry single amino acid substitutions in the central domain of NTRC (Figure lB), the domain required for its ATPase activity (Table 1 C). The ATPase activity of phosphorylated NTRC was

Cell 158

Table

2. ATPase

Activities

of NTRC’“P’““““’

P, Release

Protems

(pmol/lO

HI)

Protein

+NTRB

-NTRB

Net’

Percentage of Wild-Type ATPase Activity”

Wild type G173N

297

ND 1.6

297

100

Wild Type S207F E208Q G219K R294C R358C R358H

339 36 53 190 25 20 21

4.0 6.4 14 2.7 1.2 2.0 2.8

335 30 39 187 24 18 18

9.5

NTRC-Phosphate (pmolll0 ~1)

3

2.1 3.5

9 12 56 7 5 5

2.3 2.5 2.2 2.6 4.0 4.2 4.6

7.9 100

ATP hydrolysis (P, release) by NTRC’“P’“‘^’ proteins was measured in the presence (+NTRB) and absence (-NTRB) of phosphorylation, as described in Experimental Procedures, and is given in units of pmol/lO @I of reaction mixture. Data for the G173N protein are from the experiment in Figure 4, whereas data for the other proteins are from a separate experiment in which the specific activity of ATP was 2401 cpmlpmol, and the background of “P, was 13 pmol/lO ~1. P, release was measured 20 and 21 min after addition of label for the G173N and other mutant proteins, respectively. P, release in the absence of NTRB provides a measure of contaminating ATPase activity in the preparation and is subtracted from P, release in the presence of NTRB to obtain “net” release. Net release is a maximum estimate of ATPase activity, since P, release due to the autophosphatase activity of NTRC-phosphate has not been subtracted. The amount of NTRC-phosphate was determined from steady-state levels of P, incorporation, as descrrbed in Experimental Procedures. ND, not determined. a “Net P, release” was not corrected for P, release owing to the autophosphatase activity of NTRC-phosphate. This correction is approximated as follows for the NTRCG”IN protein during the 20 min between addition of label and sampling for P,. The half-life for NTRCG’73N-phosphate is 3 min (not shown), so one-sixth of the incorporated P, is released per minute. It takes about 10 min for the label to equilibrate into NTRC-phosphate (Figure 28). To calculate P, release during this period, we assume an average labeled pool size of 1.75 pmol/lO frl (i.e., half that after equilibration); P, release due to autophosphatase is then (1.75 pmoll6 min) x 10 = 3 pmol. During the 10 min after equilibration of label into NTRC-phosphate, P, release is (3.5 pmoll6 min) x 10 min = 6 pmol. Total P, release due to the autophosphatase is therefore *9 pmol. b Not corrected for autophosphatase activity.

grossly impaired in NTRC’BPr”s*o’proteins with the following lesions: Glyl73-Asn, Arg294+Cys, Arg358+Cys, and Arg358-His (Table 2). This activity was not so grossly defective in other NTRC’*PrBS*“’ proteins (Ser207+Phe, Glu208+Gln) and was little impaired in one of them (Gly219-Lys). The Glyl73-Asn protein, which shows the greatest defect in ATPase activity, is particularly interesting, because the lesion changes a conserved residue in the putative ATP-binding motif. The 3% residual activity reported for this protein in Table 2 is a maximum estimate, since it includes P, release by the autophosphatase. (The rate of P, release due to the autophosphatase can be inferred from the steady-state level of NTRC-phosphate, the time course of labeling, and the half-life of NTRC-phosphate. Calculations [Table 21 indicate that m9 pmol of P, would have been released by the autophosphatase at the

Table

3. Comparison

of ATPase

Actrvities

of Wild-Type

and

NTRCc”“s”‘U”“”

Unphosphorylated P, Release 60 minll0

Protein Wild type Constitutive

(S16OF)

5 43

(pmol/ VI)

20 min time point used in Table 2, so the observed 8 pmol of P, release by the mutant protein can be accounted for by the autophosphatase.) When measurements were made at early time points and hence were not complicated by release of labeled P, due to the autophosphatase activity, no Pi release by the mutant protein was detected (Figure 4). NTFiCS160F (Constitutive) Has ATPase Activity in the Absence of Phosphorylation As stated above, NTFiCS’60F has some ability to activate transcription without being phosphorylated, unlike wildtype NTRC. We therefore predicted that unphosphorylated NTRCSiwF should have more ATPase activity than unphosphorylated wild-type NTRC. To demonstrate this difference convincingly, we found it necessary to purify

Proteins Phosphorylated

Specific Activitya (pmol/mm/pmol)

P, Release (pmoll 5 min/lO frl)

Specific Activityt (pmol/min/pmol)

0.008 0.07

130 750

20 28

To determine the rate of P, release in the absence of phosphorylation, 1 PM NTRC protein was warmed in acetate buffer (Experimental Procedures) at 37% for 4 min, at which time 0.4 mM MgATP (1363 cpmlpmol) was added. Samples were taken for P, determination after 60 min. The rate of P, release by phosphorylated NTRC was determined by the standard prephosphorylation procedure, except that the buffer was acetate buffer. The specrfic activity of ATP was 2212 cpmlpmol. The background of P, was 30 pmol. d One mrcromolar NTRC is equrvalent to 10 pmol dimerll0 PI of reaction mixture. : ’ The steady-state levels of NTRC-phosphate (pmol of P, incorporated) were 1.3 pmol/lO ~1 and 5.3 pmol/lO frl for the wild-type and constitutive proteins, respectively. The specific activrties are a minimum estimate, in that the rate of ATP hydrolysis is not linear with NTRC-phosphate concentratron at the levels of NTRC-phosphate present in this experiment (Figure 6A). In particular, the activity of the constitutive protein is underestimated

Phosphorylated 159

NTRC

Is an ATPase

A

Figure

B

3. Nucleotide

Specificity

of the ATPase

NTRC-phosphate was generated by preincu1000 400. bating 1 FM NTRCS’“F (constitutive) and 100 nM NTRB with 0.4 mM unlabeled ATP for 10 a. min. The reaction mixture was then split into several tubes, each of which contained a carrier-free “P-labeled nucleoside triphosphate. For the experiment in (A), the nucleotides used were v-labeled ATP (open squares, 1525 cpm/pmol), a-labeled ATP (filled squares, 220 cpm/pmol), a-labeled CTP (filled triangles, 170 cpmlpmol ATP). and a-labeled UTP (filled triangles, 150 cpmlpmol ATP). For the experiA ment in (B), the nucleotides used were y-labeled ATP (open squares, 2700 cpmlpmol) and 0 20 y-labeled GTP (open triangles, 2300 cpmlpmol 40 60 80 0 10 20 30 40 50 ATP). Samples were taken at the times indiTime (min) Time (min) cated and analyzed for release of V or 3*Pnucleoside diphosphate by thin-layer chromatography as described in Experimental Procedures. The backgrounds of endogenous V were 100 pmol/lO ul for y-labeled ATP in (A) and 4 pmol/lO ul for v-labeled ATP and GTP in (6). The backgrounds of endogenous a3*P nucleoside diphosphate were 16, IO, and 85 pmol/lO ul for ATP, CTP, and UTP, respectively. (y-Labeled nucleotides were used at higher specific activity than a-labeled nucleotides to facilitate detection of phosphorylated NTRC.) The steady-state levels of NTRC-phosphate were 1.4 pmol/lO PI and 1.2 pmolll0 trl for (A) and (6) respectively

the proteins more highly to remove contaminating ATPase activities. These contaminating activities differed in the two preparations, because NTRCS’GOF eluted from heparin-agarose at a higher salt concentration than wild-type NTRC. Contaminants could be removed either by chromatography on single-stranded DNA or Mono-Q (not shown); we did both. After purification in this fashion, unphosphorylated NTRCS’WF had about 10 times higher ATPase activity than unphosphorylated wild-type NTRC (Table 3). The ATPase activity of both proteins could still be activated by phos-

phorylation. To confirm that the activity of the NTRCSIWF protein in the absence of phosphorylation was inherent in this protein, we showed that ATPase activity cochromatographed with the protein over both a Mono-Q column (Figure 5) and a sieving column (Superose 12; not shown), as did the phosphorylation-dependent activity. NTRCStGOF(Constitutive) Has an Enhanced ATPase Activity To compare the ATPase activity of phosphorylated NTRCS”joF (constitutive) to that of wild-type NTRC, we 80

0.8

60 -8 1 h %40 is c g '20

0.6

58

60

62 Fraction

Figure

0

10

20

30

40

Time (min) Figure

4. The ATPase

Activity

of NTRCG’73N

(Repressor)

The time course of ATP hydrolysis by phosphorylated wild-type NTRC (open squares) or the repressor protein NTRCGi73N with phosphorylation (filled squares) and without phosphorylation (filled circles) was determined as described in Experimental Procedures. Each point on the graph is the mean of duplicate reactions. Error bars indicate the standard deviation for cases in which it is larger than the symbol. The specific activity of ATP was 2912 cpmlpmol, the Y background was 12 pmol, and the steady-state levels of NTRC-phosphate in the presence of NTRB were 2.1 and 3.5 pmol/lO trl for wild-type and G173N proteins, respectively.

5. Coelution

of ATPase

Activity

64

66

68

70

Number with NTRCS’MF

(Constitutive)

Purified NTRCS’MF was chromatographed on a high resolution anion exchange column (Mono-Q) as described in Experimental Procedures, The eluted fractions were assayed for protein content (open diamonds) and for ATPase activity in the presence of NTRB (filled squares, “phosphorylated”) and the absence of NTRB (filled circles, “unphosphorylated”). ATPase assays were performed at 1 FM NTRC, except that fraction 60, which had very little protein, was assayed at 0.1 PM. Samples for P, release were taken at 5 and 15 min after addition of label rn the presence and absence of NTRB, respectively. The specific activity of ATP was 2212 cpmlpmol in the presence of NTRB and 1363 cpml pmol in its absence. The background of “P, was 30 pmol in each case. The buffer in this assay contained acetate rather than chloride as amon; acetate was found to be stimulatory and facilitated detection of ATP hydrolysis by the unphosphorylated S16OF protem.

Cell 160

-a-

5-fold in two additional experiments (not shown). The greater ATPase activity of phosphorylated NTRCS’60F made it the preferred protein for assays in which sensitivity of detection was an issue. We could not determine a specific activity for the ATPase of phosphorylated NTRC, because ATPase activity was not linear with NTRC-phosphate. For both wild-type NTRC and NTRCSIWF, the ATPase activity increased with increasing NTRC-phosphate, but the relationship was complex (Figure 6A). Activation of the ATPase appeared to require a threshold concentration of NTRC-phosphate. Above the threshold, activity increased dramatically, but then abruptly reached a plateau. The unusual dependence of activity on NTRC-phosphate concentration will be discussed below.

NTRCwild-type

-NTR@'~O~

0

2 1 NTRC-phosphate

3 (pmoV10

4 ~1)

^

1 0 %

loo

s E

0 0

Figure 6. Dependence NTRC-Phosphate

1 2 NTRC-phosphate of the ATPase

3

4

(pmoV10 Activity

5

ul)

on the Concentration

of

(A) Comparison of wild-type NTRC (open squares) with NTFKYMF (filled squares). To generate different amounts of NTRC-phosphate, 1 uM NTRC was incubated with 0.4 mM unlabeled ATP and increasing amounts of NTRB (0, 1, 24, 6, 12.5, 25, 50, 75, and 100 nM) at 37V for 10 min, at which time [rJZP]ATP was added. The final specific activity was 1470 cpmlpmol, and the ? background was 17 pmol/lO ul of reaction mix). Samples were taken at 5 min to determine the rate of ATP hydrolysis and at 15-25 min to determine the level of NTRC-phosphate, as described in Experimental Procedures. (6) The effect of polyethylene glycol and glycerol on the ATPase activity of phosphorylated wild-type NTRC. Conditions as in (A), except that one set of reaction mixtures contained 3.5% polyethylene glycol8000 (open circles), and one contained 10% glycerol (open diamonds). The NTRS concentrations used were 0, 1,255, 10, 25, 50, 100, 200, and 300 nM. The specific activity of ATP was 2103 cpmlpmol, and the 32P, background was 15 pmol/lO ml of reaction mix.

phosphorylated the two proteins to various extents and compared their ability to release P, from [yJ*P]ATP. The proteins were phosphorylated by adding different amounts of NTRB (O-100 nM) to a fixed amount of NTRC in the presence of unlabeled ATP and incubating for 10 min. After this, [yJ2P]ATP was added to measure ATPase activity (P, release) and the steady-state level of NTRCphosphate (incorporation of P, into protein). Phosphorylated NTRCSIGOFwas more active than wild type by two criteria: less phosphate had to be incorporated to activate the ATPase, and the maximum rate of hydrolysis was greater (Figure 6A). The difference in maximum rate was 3-fold in the experiment shown in Figure 6A and 4- and

The Nucleotide Specificity of the ATPase Is the Same As That for Open Complex Formation As stated above, NTRCSJmF (constitutive) requires ATP to catalyze open complex formation, even in the absence of NTRB. Studies of the nucleotide specificity for open complex formation showed that GTP could substitute for ATP, albeit weakly, but CTP and UTP could not (Popham et al., 1989). We found that the ATPase exhibits the same specificity. Phosphorylated NTRCSlmF hydrolyzed GTP at about one-tenth the rate of ATP (Figure 3B), consistent with the weak ability of GTP to support open complex formation. The GTPase activity could not be attributed to contamination of the GTP with ATP at a level of IO%, because visual inspection of thin-layer chromatograms did not reveal any ATP (not shown). The time course of Pi release from [yJ2P]GTP, which was fastest at the start of the assay, together with the fact that there was no detectable incorporation of P, into NTRC (not shown), indicated that the observed hydrolysis of GTP was not due to the autophosphatase activity. Phosphorylated NTRCSlwF did not detectably hydrolyze CTP or UTP (Figure 3A), consistent with the inability of these nucleotides to support open complex formation. ATPyS and ADP inhibit Both the Transcriptional Activation and ATPase Activities of NTRC Although we knew that ADP and a number of analogs of ATP with a nonhydrolyzable 8-r bond could not support open complex formation (Popham et al., 1989) we did not know whether this was due to their failure to be hydrolyzed or their failure to bind to NTRC. To determine whether any of these analogs could in fact bind to NTRC, we determined whether they could inhibit catalysis of open complex formation by NTRCSIGOFin the presence of ATP. Titrations revealed that ATPyS (adenosine-5’-O-[3-thiotriphosphate]) and ADP are potent inhibitors of open complex formation in the presence of ATP (Figure 7A). When ATP was present at 1 mM, 0.25 mM ATPyS or 0.4 mM ADP reduced open complex formation by half; 2 mM ATPyS or 3 mM ADP completelyprevented open complex formation. High concentrations of ATP restored open complex formation in the presence of ATP$S (Figure 78). We considered the possibility that ATPyS and ADP did in fact support open

Phosphorylated 161

NTRC

Is an ATPase

A

1.0 Inhibitor

-

ATpVS

-+-

ADP

complex formation but prevented transcription because ATP hydrolysis was needed to dissociate es4 holoenzyme from NTRC. This possibility was excluded by the observation that ATPyS and ADP inhibited open complex formation, as assessed by a gel mobility shift assay (not shown); open complexes, but not closed complexes, retard the mobility of a DNA fragment bearing the g/nA promoter (Popham et al., 1989). ATPyS and ADP were also found to be potent inhibitors of the ATPase activity of phosphorylated NTRCS’GoF; when the ATP concentration was 0.4 mM, approximately0.2 mM ATPyS or 0.35 mM ADP reduced activity by 50% (Figure 7C). For this assay, P, release was determined 1 min after addition of the inhibitor to phosphorylated NTRC to ensure that comparable levels of NTRC-phosphate were present throughout the range of inhibitor concentrations tested (both ATPyS and ADP inhibited phosphorylation). Control experiments indicated that phosphorylated NTRCS’GOFhydrolyzed [y@S]ATPyS at l%-2% the rate of ATP (not shown); unphosphorylated protein did not detectably hydrolyze ATPrS.

2.0

(mM)

-0-OmhtATPyS -.z- 0.4mMATPyS + 0.6mMATPyS

-0.0

2.0

ATP

t

0.0

0.5

1.0 Inhibitor

Figure ATPyS

7. Inhibition and ADP

of Open

Complex

6.0

4.0

(mM)

ADP

1.5

2.0

(n&f) Formation

and

ATPase

by

(A) and (B) show inhibition of open complex formation, and (C) shows inhibition of ATPase. (A) NTRCS’QF (constitutive) was used in the absence of NTRB to catalyze formation of open complexes by as4 holoenzyme at the g/nA promoter, as described in Experimental Procedures. Open complexes were formed in the presence of 1 mM ATP and the indicated amount of ATPyS (open squares) or ADP (filled diamonds) and were quantitated by their ability to yield transcripts in a single-cycle transcription assay. Control experiments indicated that neither analog inhibited initiation or elongation of transcripts once open complexes had been formed (not shown). (B) As in (A), except that open complexes were formed in the presence of 0 (open circles), 0.4 (open squares), or 0.6 (filled squares) mM ATPyS and the indicated amount of ATP. (C) NTRC-phosphate was generated by preincubating 1 uM NTRCSIMF and 100 nM NTRB with 0.4 mM unlabeled ATP for 10 min. The reaction was then split into several tubes, each of which contained carrier-free [y-32P]ATP and enough ATPyS (open squares) or ADP (filled diamonds) to achieve the indicated final concentration. Samples for P, release were taken 1 min after addition of label. The level of NTRC-phosphate at steady state with ATP alone was 5.6 pmol/lO ul of reaction mix. Although the analogs inhibit phosphorylation, the level of NTRCphosphate changes by less than 10% during the 1 min incubation, because the half-life of the phosphorylated S16OF protein is about 5 min (J. Keener and S. Kustu, unpublished data).

Activation of the ATPase May Involve Cooperative Interactions among Dimers of NTRC-Phosphate Our initial investigations of the unusual dependence of ATPase activity on NTRC-phosphate concentration (Figure 6A) indicated that it was not an artifact of the mode of assay. Essentially the same activity curve was obtained when the amount of NTRC-phosphate was varied by three alternative protocols: holding NTRC fixed at a high concentration and titrating NTRB, as described above (Figure 6A); holding NTRB fixed at a high concentration and titrating NTRC; and diluting phosphorylation reactions to various extents (not shown). We interpret this to mean that the concentration of NTRC-phosphate, rather than the ratio of NTRC to NTRB or the absolute amount of proteins in the assay, is the important variable. These results argue against the idea that the activity plateau seen in Figure 6A occurs because high concentrations of NTRB are inhibitory. We considered three trivial explanations for the observation that the ATPase activity reaches a plateau with increasing NTRC-phosphate: limitation for ATP, inhibition by ADP or P,, and the presence of a large fraction of inactive NTRC molecules in NTRC preparations. We doubt that ATP is limiting at the plateau, because P, is measured after 5 min, when <5% of the ATP has been consumed (~10% in the case of NTRCSiGoF). Moreover, the rate of ATP hydrolysis is roughly linear for about 20 min. The linearity with time also argues against inhibition by ADP or P, being responsible for the plateau. Although we do not know what fraction of our NTRC is active, we note that titrations performed with two independent preparations of wild-type NTRC yielded nearly superimposable activity curves (not shown). Although a requirement to phosphorylate both subunits of NTRC, which is a dimer in solution, could contribute to the complex activity curve, we do not think such a requirement could account for it. First, the rise is too steep: if subunits are phosphorylated independently, the ATPase

Cell 162

activity would be expected to increase linearly with the square of the NTRC-phosphate concentration. In our titrations, activity appears to be proportional to approximately the third to fourth power of the NTRC-phosphate concentration. Second, the plateau is reached at a low concentration of NTRC-phosphate; for example, the ATPase activity of wild-type NTRC reaches a plateau when only 1 pmol of phosphate has been incorporated per 10 ul of reaction mixture, which corresponds to filling of only ~5% of the phosphorylation sites (Figure 6A). Our working hypothesis to account for the complex relationship between the concentration of NTRC-phosphate and the rate of ATP hydrolysis is that the aggregation state of NTRC-phosphate affects the ATPase activity: the threshold reflects a stimulation of the ATPase when phosphorylated dimers of NTRC interact, whereas the plateau reflects a solubility limit for phosphorylated NTRC. This hypothesis is based on two observations. Phosphorylated NTRC has a demonstrable tendency to aggregate, and agents that increase or decrease aggregation affect the ATPase activity in a manner consistent with the hypothesis. When present at high concentration (4 PM), wild-type NTRC forms a visible precipitate upon phosphorylation, and NTRCSIGoFdoes so even in the absence of phosphorylation. Although turbidity is not obvious during a standard ATPase assay, in which the concentration of NTRC is 1 PM, monitoring of light scattering at 350 nm indicated that aggregation occurred and was stimulated lOO-fold by phosphorylation (not shown). Elevation of the glycerol concentration from 3% to 10% greatly inhibited aggregation, as assessed by light scattering at 350 nm, and correspondingly shifted the ATPase activity curve to higher concentrations of NTRC-phosphate (Figure 6B). Addition of polyethylene glycol (final concentration 3.5% [w/v]), which is known to increase effective macromolecular concentrations (Minton, 1983) shifted the activity curve to lower NTRC-phosphate concentrations and decreased the maximum activity (Figure 6B). Discussion NTRC Has a Required ATPase Activity The enhancer-binding protein NTRC catalyzes open complex formation by o 54 holoenzyme in a reaction that requires ATP (Popham et al., 1989). To catalyze open complex formation, NTRC must be phosphorylated (Ninfa and Magasanik, 1986). In this paper we have demonstrated that the phosphorylated form of NTRC hydrolyzes ATP to ADP and P,. This ATPase activity appears to be related to the ATP requirement for open complex formation, because both reactions have the same nucleotide specificity (Figure 3). More convincing, some mutant forms of NTRC that fail to activate transcription (NTRC’ePre*Sor)have an impaired ATPase activity (Figure 4 and Table 2). In particular, a change of a conserved residue in the putative ATPbinding motif of NTRC (Glyl73-Asn in the central “activation” domain of the protein) resulted in a protein with little or no ATPase activity. The existence of a point mutation that simultaneously deprives NTRC of its ability to activate transcription and hydrolyze ATP argues strongly that NTRC must interact with ATP to catalyze open complex

formation. However, the finding that ATP hydrolysis is impaired in NTRCr*PrBsso’proteins does not distinguish a requirement for ATP hydrolysis from a requirement for ATP binding, because nothing is known about the ATP-binding properties of the mutant proteins. If ATP hydrolysis is required for open complex formation-i.e., if ATP binding is not sufficient-we would make two predictions, both of which have been fulfilled. First, any NTRC protein that is capable of catalyzing open complex formation should have ATPase activity. We have demonstrated that phosphorylated NTRC has an ATPase activity. In addition, we have demonstrated that an NTRCconstitUtlve protein (SerlGO-Phe) that has the ability to catalyze formation of open complexes without being phosphorylated also has detectable ATPase activity in the absence of phosphorylation (Figure 5 and Table 3). The ATPase activity of NTRCSiBOF is ~1 O-fold greater than that of unmodified wild-type NTRC, which cannot catalyze open complex formation. Second, if there are nonhydrolyzable analogs of ATP that fail to support open complex formation but retain the ability to bind to NTRC (NTRC-phosphate), they should block both ATP hydrolysis and the ability of ATP to support open complex formation. ATPyS and ADP are such analogs (Figure 7). Since ATP$S does not support open complex formation, we infer that binding of ATP is not sufficient to allow NTRC-phosphate to activate transcription. Since ADP also fails to support open complex formation, ATP hydrolysis does not function to change the conformation of NTRCphosphate from an inactive ATP-bound form to an active ADP-bound form. Rather, the hydrolysis event itself appears to be required. Presumably, ATP hydrolysis is supplying energy to overcome an activation energy barrier to open complex formation or to make this change in configuration of the polymerase thermodynamically favorable. Among the mutant forms of NTRC (NTRCrePrBSsor)that fail specifically in transcriptional activation, at least one protein (Glyl73+Asn) has so little ATPase activity that its failure to catalyze open complex formation can probably be attributed to lack of ATP hydrolysis per se. Although the other NTRCrePressorproteins have a defective ATPase, we note that most appear to have more activity than unphosphorylated NTRCS’“OF, which can catalyze open complex formation. The most striking example is the NTRCepressor protein Gly219-+Lys, which has nearly wildtype ATPase activity and yet has no detectable ability to activate transcription. It seems unlikely that diminished ATPase activity accounts for the profound inability of this protein to activate transcription. The primary defect of this protein may lie in coupling of ATP hydrolysis to open complex formation or in protein-protein interaction with os4 holoenzyme. The latter appears to be required for DNA loop formation between NTRC and polymerase (Su et al., 1990) and would presumably be another function required of the central domain of NTRC. The ATPase Activity of NTRC Is Regulated by Phosphorylation Phosphorylation of the amino-terminal regulatory domain of NTRC stimulates by at least several hundred fold the

Phosphorylated

NTRC Is an ATPase

163

ability of NTRC to hydrolyze ATP (Tables 1 and 3). We were unable to find stimulation of the ATPase by two potential effecters other than phosphorylation-DNA and o54 holoenzyme (not shown). We are confident that neither DNA nor CS~ holoenzyme activates ATP hydrolysis by unphosphorylated wild-type NTRC. However, owing to the coupled nature of the ATPase assay and the complex dependence of ATPase activity on NTRC-phosphate concentration (Figure 6), we cannot rule out the possibility that these effecters might stimulate the ATPase activity of NTRC-phosphate under appropriate conditions. Our finding that the ATPase activity of wild-type NTRC is dependent upon phosphorylation plausibly accounts for the observation that NTRC must be phosphorylated to function as a transcriptional activator (Ninfa and Magasanik, 1986). Phosphorylation may play roles in addition to its apparently essential role in activating the ATPase. Although phosphorylation is not required for NTRC to bind with high affinity to its enhancer-like sites upstream of g/nA, phosphorylation increases the cooperativity of binding (S. Porter and S. Kustu, unpublished data). Phosphorylation appears to be required in vivo for cooperative binding of NTRC to sites upstream of the &dependent niflA promoter of Klebsiella pneumoniae (Minchin et al., 1988); these sites are much weaker than those at glnA (Wong et al., 1987). Activation of the ATPase activity of NTRC is a defined mechanism by which phosphorylation regulates the function of a receiver protein in a two-component regulatory system. (Receiver proteins have a domain homologous to the amino-terminal regulatorydomainof NTRC [Figure 11.) We note parenthetically that activation of an ATPase is likely to pertain only for receiver proteins that share the central “activation” domain of NTRC (see below). All Activators of 0% Holoenzyme Probably Have an ATPase Activity The ATPase activity of NTRC appears to be associated with an ATP-binding motif in its central domain (Tables 1 and 2). Consistent with this, changing the first serine in the ATP-binding motif to alanine results in an NTRC protein that cannot activate transcription but still binds to DNA (Drummond et al., 1990). Every activator of as4 holoenzyme whose sequence is known contains a domain homologous to the central domain of NTRC (Kustu et al., 1989; Thony and Hennecke, 1989). Since the ATP-binding motif is among the most highly conserved features of this domain, we predict that all activators of 05“ holoenzyme will have an ATPase activity under appropriate conditions and that ATP hydrolysis will be required for their function. In support of this, a lesion homologous to the Glyl73-Asn change in the ATP-binding motif of NTRC abolishes transcriptional activation by HRPS, an activator of @‘-dependent genes whose products are required for virulence of the plant pathogen Pseudomonas syringae, pathovar phaseolicola (R. Fellay and N. Panopoulos, personal communication). Although we expect all activators of os4 holoenzyme to have a required ATPase activity, we expect this activity to be regulated by phosphorylation only in those activators that are also receiver proteins in two-component regula-

tory systems-ALGB, DCTD, FLED, and HOXA (Ronson et al., 1987; Ramakrishnan and Newton, 1990; Eberz and Friedrich, 1991; Wozniak and Ohman, 1991). In the case of activator proteins that do not have a domain homologous to the amino-terminal regulatory domain of NTRCFHLA, HRPS, NIFA, and XYLR (Buikema et al., 1985; Drummond et al., 1986; lnouye et al., 1988; Grimm and Panopoulos, 1989; Maupin and Shanmugam, 1990; Schlensog and Bock, 1990)-it appears likely that ATPase activity will be constitutive or will be regulated by mechanisms other than phosphorylation. Requirements for ATP Prior to Initiation of Transcription in Other Systems An ATP requirement prior to initiation of transcription is rare in bacteria. We are aware of only two examples other than activation of os4 holoenzyme. The activator of T4 late transcription catalyzes the formation of open complexes by an alternative holoenzyme form of RNA polymerase containing two TCencoded proteins (Kassavetis et al., 1983; Herendeen et al., 1989, 1990). Open complex formation is dependent on ATP hydrolysis by an enhancerbinding protein (Herendeen et al., 1989). The enhancerbinding protein has a DNA-dependent ATPase activity and also functions as a “sliding clamp” that increases the processivity of DNA replication. The maltoseT (MALT) protein of Escherichia coli stimulates formation of open complexes by 070 holoenzyme. To bind DNA and activate transcription, MALT requires ATP and maltotriose, both of which bind to MALT with high affinity (Richet and Raibaud, 1989). Although MALT is capable of ATP hydrolysis and hydrolysis is stimulated 2-to 3-fold by maltotriose, nonhydrolyzable analogs of ATP can support activation of transcription by MALT. Thus, ATPbinding appears to be sufficient for the function of MALT as a transcriptional activator. In contrast to the situation in bacteria, an ATP requirement prior to initiation of transcription is common in eukaryotes. ATP is required for the formation of a preinitiation complex by RNA polymerase III (C), the assembly of which is the rate-limiting step in 5S RNA gene transcription (Bieker et al., 1985). Since nonhydrolyzable analogs of ATP can substitute in this reaction, ATP binding appears to be sufficient (Bieker and Roeder, 1986). ATP hydrolysis is required for initiation of transcription by mitochondrial polymerase (Narasimhan and Attardi, 1987) and is also required prior to or concomitant with synthesis of the first phosphodiester bond by RNA polymerase II (B) (reviewed in Sawadogo and Sentenac, 1990; see also Luse and Jacob, 1987). Finally, initiation of transcription of vaccinia virus early (Shuman et al., 1980) and intermediate (Vos et al., 1991) genes by viral RNA polymerases depends on ATP hydrolysis. ATPases are being characterized in the latter systems (Broyles and Moss, 1988; Conaway and Conaway, estingly,

1989; Soptaet al., 1989; Voset al., 1991); interthey apppear to be general transcription factors

rather than enhancer-binding Experimental Construction All enzymes

proteins.

Procedures of Plasmids

for Protein

used lo manipulate

Overproduction

DNA were purchased

from New En-

Cell 164

gland BioLabs or Boehringer Mannheim. DNA isolation and cloning were carried out following standard procedures. The vectors used for expression of proteins were pT7-5 and pT7-7 (Tabor, 1990), both of which allow transcription of inserted DNA fragments from the 010 promoter of phage T7 (dependent on T7 RNA polymerase). Plasmid pT7-7 carries, in addition, the ribosome-binding site for the ~10 gene and an open reading frame in its polylinker. Plasmids for the Overproduction of NTRC Proteins Plasmids that allow high levels of overproduction of NTRC (-10% of total protein; not shown) were constructed in the following steps. First, a 1.9 kb EcoRI-Sail fragment from pJES222 (D. Weiss and S. Kustu, unpublished data) carrying the complete Salmonella typhimurium ntrC gene and 87 bp of upstream DNAwas inserted into pT7-7 that had been digested with EcoRl and Sall. Second, this construct was digested with Ndel to linearize it at the ATG of the open reading frame within the polylinker and PflMI, which cuts at a unique site in codons 15 and 16 of ntfC. The T7 ribosome-binding site was joined to ntrC with a double-stranded synthetic oligonucleotide (47/44-mer) that begins with the ntrC ATG and continues into ntrC sequences to the PflMl site. (This oligonucleotide preserves the amino acid sequence of NTRC but encodes a Sall site not originally present in the gene.) The ntfC gene in this plasmid directed synthesis of a truncated NTRC protein, presumably because a frameshift mutation had occurred during one of the cloning steps. Third, the frameshift was corrected by digesting with EcoRV and Hindll to delete most of ntrC and inserting an EcoRVHindll fragment from pJES46 (Hirschman and Kustu, unpublished data) to reconstruct the gene. We call the resulting plasmid pJES297. It overproduces NTRC to >lO% of cell protein, but it inhibits growth of an E. coli host even in the absence of T7 RNA polymerase (not shown). We suspected that the growth defect resulted from transcription of r&C by a host RNA polymerase initiating at an unidentified promoter(s) in the vector. Fourth, plasmid pJES297 was digested with BstBI, which cuts 20 and 30 nucleotides upstream of the transcriptional start site for the ~10 promoter; and the T7 early terminator, a strong transcriptional terminator of bacterial RNA polymerase, was inserted on an ~300 bp HinPl fragment from pTElO2 (Elliott and Geiduschek, 1984) to create plasmid pJES311. The terminator in pJES311 is in the “wrong”orientation. Although both orientations of the terminator alleviated the growth defect in the absence of T7 RNA polymerase, synthesis of ntrC could not be induced when the terminator was in the “correct” orientation. We do not know the reason for this. To overproduce mutant forms of NTRC, various ntrC alleles were subcloned into pJES311 by replacing the EcoRV-EcoRI fragment of pJES311 with a corresponding fragment from other plasmids. This replaces all of NTRC downstream of codon 54. The following plasmids were constructed in this manner: pJES392 (ntrC635, Gln173-Asn); pJES393 (ntrC638, Arg358-Cys); pJES395 (ntrC640, Gly219-Lys); pJES396 (ntrC647, Arg358-His); pJES397 (ntrC642, Glu208-Gin). Because the ntrC73 mutation creates an EcoRl site, pJES412 (ntrC73, Ser207+Phe) was constructed by replacing the EcoRV-Hindlll fragment of pJES311 with a 1.8 kb EcoRV fragment. This creates an illegitimate EcoRV-Hindlll joint that lies outside (downstream) of ntrC. Two mutant NTRC proteins were overproduced under control of the leftward promoter of phage 1 in plasmid PLc28 (Remaut et al., 1981). The relevant plasmids are pJES76 (ntrC670, SerlGO-Phe) and pJES97 (ntrC72, Arg294-Cys).

Plasmid for the Overproduction of NTRB Plasmid pJES278. which allows high levels of overproduction of NTRB from S. typhimurium (~10% of cell protein; not shown) was constructed from pJES23 (J. Hirschman and S. Kustu, unpublished data) in the following steps. First, a 2.9 kb Xholl-Hindll fragment carrying most of ntr6 (Xholl cuts at codon 12) and all of ntrC was ligated into pT7-5 that had been digested with BamHl and Hindlll. Ligation of the BamHl end to the Xholl end recreated the BamHl site. Second, most of the ntrC gene was deleted by digesting with EcoRV, which cuts in codon 54, and Hindlll, which cuts downstream of ntrC, treating with Klenow fragment to make the Hindll end blunt, and religating. Third, the complete ntrB gene was reconstructed by digesting with EcoRI, which cuts in the polylinker of pT7-5, and BamHI, which cuts at the juncture of the vector and ‘ntr6, and ligating in a double-stranded

synthetic site and

oligonucleotide(54/54-mer) encodes the first 12 amino

that contains a ribosome-binding acids of NTRB.

Overproduction of Proteins Plasmids for overproduction of NTRC proteins could not be maintained in E. coli strains that contained T7 RNA polymerase; therefore, plasmids were transformed into an E. coli Hfr strain, and induction was accomplished by infecting with an Ml3 phage that carries T7 RNA polymerase under control of the lac promoter. The strain used for overproduction of NTRC proteins was NCM724 (HfrC[;1], WAG, argG:: TnlO], a derivative of E. coli K38 (Tabor, 1990). Cultures were grown in maximal induction medium (Mott et al., 1985) containing 2 mM glutamine and 200 pglml ampicillin at 37OC with vigorous shaking. At an OD,, of ~1, shaking was slowed to prevent shearing of pili, and T7 RNA polymerase was introduced by adding Ml 3 mGPl-2 to a multiplicity of infection of about 10 (Tabor, 1990). Isopropyl-g-D-thiogalactoside (IPTG) was also added to 0.5 mM to allow induction of T7 RNA polymerase. After 2 hr cultures were harvested by centrifugation. The plasmid for overproduction of NTRB (pJES278) could be stably maintained in E. coli in the presence of T7 RNA polymerase; the host strain used was NCM699 (AlacTnlO hsdSga/ 1DE3: /ac//acU&gene 1 [T7 RNA polymerase]), which is derived from BL21 (1DE3) (Studier and Moffatt, 1986). NCM699 carries two plasmids that reduce transcription by T7 RNA polymerase prior to induction: pJES296, an F’that contains laclq; and pLysE, which expresses T7 lysozyme, an inhibitor of T7 RNA polymerase (Studier et al., 1990). To overproduce NTRB, strain NCM703 (NCM699 carrying pJES278) was grown in maximal induction medium with 200 bglml ampicillin and 10 pglml chloramphenicol at 37OC with vigorous shaking. At an OD, of -1, IPTG was added to a final concentration of 0.5 mM to induce T7 RNA polymerase and thereby NTRB. After 45 min, rifampicin was added to 50 Bglml to reduce the expression of E. coli proteins under control of E. coli RNA polymerases. Cells were harvested by centrifugation after an additional 75 min. Purification of Proteins All procedures were performed at 4OC in a standard buffer that contained 10 mM Tris titrated to pH 8.0 with acetic acid, 50 mM KCI, 0.1 mM EDTA, 5% glycerol, and 1 mM dithiothreitol (DTT). Protein concentrations were determined by the Bradford method (Bio-Rad) with bovine serum albumin (BSA) as standard. NTRC and NTRB are dimers in solution. NTRC was purified essentially as described (Hirschman et al., 1985). Briefly, NTRC was precipitated with ammonium sulfate (O%35%) and was then chromatographed on Q-Sepharose Fast-Flow (Pharmacia), heparin-agarose (Bethesda Research Labs), and Mono-Q (Pharmacia). The purified protein was >95% pure as judged by silver staining of SDS-polyacrylamide gels. For some experiments NTRC was further purified by chromatography on single-stranded DNA-agarose (Bethesda Research Labs) to remove contaminating DNA-dependent ATPases, which eluted later than NTRC. NTRC protein in the standard purification buffer was applied to a 10 ml single-stranded DNAcolumn that had been equilibrated with the same buffer. Wild-type NTRC eluted isocratically and was subsequently concentrated bychromatographyon Mono-Q. NTRCSIMF was eluted with a 150 ml linear gradient from 50 to 500 mM KCI. The S16OF protein eluted at about 150 mM KCI. The flow rate was 0.5 ml/ min for all steps, and 4 ml fractions were collected. To determine whether ATPase activity coeluted with NTRCSIMF from Mono-Q, 1.6 mg of highly purified NTRCS’MF (through single-stranded DNA step), was loaded on a 1 ml Mono-Q column. After washing with 5 ml of buffer, NTRC was eluted with a 15 ml linear gradient from 50 to 500 mM KCI at a flow rate of 0.5 mllmin, and 0.25 ml fractions were collected. NTRCSIMF eluted at about 275 mM KCI. NTRB was purified essentially as previously described (Keener and Kustu, 1988). Briefly, NTRB was precipitated with ammonium sulfate (O%-45%) and was then chromatographed on Q-Sepharose Fast-Flow (Pharmacia), phenyl-agarose (Bethesda Research Labs), Mono-Q (Pharmacia), and Sephacryl S-200. The purified protein was >95% pure as judged-by silver staining of SDS-polyacrylamide gels. ATPase Assay A standard ATPase

assay

contained

50 mM Tris-acetate

(pH i3.0), 40

Phosphorylated 165

NTRC

Is an ATPase

mM potassium chloride, 5.4 mM magnesium chloride, 0.1 mM EDTA, 3% glycerol, 1 mM DTT, and 0.1 mglml acetylated BSA. Concentrated ATP was made equimolar with MgCI, and was then diluted to 4 mM. Carrier-free [y-32P]ATP (>5000 Cilmmol; Amersham or New England Nuclear) was diluted to 3.0 nCi/fu with water. After addition of NTRC and NTRBtothe buffer(final concentrations 1 mM and 100 nM, respectively), the mixture was warmed for 4 min at 37X, and the reaction was then initiated by adding .I vol of the stock MgATP. After allowing 10 min for the concentration of NTRC-phosphate to reach steady state, .I vol of the diluted [y-32P]ATP was added (final specific activity -2000 cpmlpmol). After 5 min, a IO nl sample for measuring P, was withdrawn and added to 990 ul of ice-cold 1 N formic acid. At 15, 20, and 25 min after addition of the label, IO nl samples for measuring NTRCphosphate were withdrawn and spotted onto Whatman ET31 filters (2 cm square), which were washed with trichloroacetic acid (TCA) and scintillation-counted as previously described (Corbin and Reimann, 1974; Keener and Kustu, 1988). The steady-state level of NTRCphosphate reported is the average of these three measurements. For some experiments (Table 3 and Figure 5) we used an acetatebased buffer that is closer in composition to our transcription buffer and that stimulated ATPase activity. This buffer is 75 mM Tris-acetate (pH 8.0) 100 mM potassium acetate, 25 mM potassium chloride, 27 mM ammonium acetate, 8 mM magnesium acetate, 0.025 mM EDTA, 2.5% glycerol, 1.5 mM DTT, and 0.1 mglml acetylated BSA. Free P, was separated from nucleoside mono-, di-, and triphosphates on polyethyleneimine (PEI) cellulose plates (Brinkman or J. T. Baker) by ascending chromatography using 0.4 M K,HPO,, 0.7 M boric acid (Bochner and Ames, 1982). Running the plates first in water improved performance. The identity of P, was determined by cochromatography with 32P, (Amersham). Samples of 5 nl were spotted onto chromatography plates, which were air dried, soaked in methanol for 5 min, air dried again, and then run. After chromatography the plates were air dried and autoradiographed. The film was used as a guide to cut out the phosphate spots from the plastic-backed TLC plates for counting radioactivity. Values reported for P, release (and NTRCphosphate) were corrected for background by subtracting the corresponding values from mock reactions that contained no proteins. To assay for hydrolysis of a-labeled nucleoside triphosphates, the following modifications were made. The labeled nucleotide was added in .l vol to achieve a specific activity of about 200 cpmlpmol of ATP. At the times indicated, 10 nl samples for determination of nucleoside diphosphate (NDP) were withdrawn and added to 90 ul of 1 .l N formic acid containing 4 mM EDTA and the corresponding NTP and NDP at 4 mM each as carriers. NDP was separated from nucleoside mono-and triphosphates by ascending chromatography on PEI cellulose plates using 0.5 M LiCI, 1 M formate (Richet and Raibaud, 1989). The identity of labeled nucleotides was determined by cochromatography with standards and detection with shortwave UV light using PEI plates with a UV,,, fluorescent background (Brinkman). To assay for hydrolysis of ATPTS, NTRCs’MFwas prephosphorylated with unlabeled ATP for 10 min in the acetate-based ATPase buffer. Then .I vol of [y-32P]ATP or [y-%]ATPyS (New England Nuclear) was added. At 5, 30, and 90 min after addition of label, 10 nl samples were withdrawn and added to 990 nl of formate that contained 1 mM cold thiophosphate as carrier and chromatographed as for P,. The identity of thiophosphate was determined by cochromatography and detection with Ellman’s reagent (Sigma; Deakin et al., 1963). Hydrolysis was quantitated by determining the ratio of P, or thiophosphate to ATP or ATPyS using a phosphorimager (Molecular Dynamics). This mode of analysis compensated for pipetting errors. To assay for inhibition of ATP hydrolysis by ATPyS or ADP, NTRCSIMF was prephosphorylated with unlabeled ATP for 10 min. Aliquots of the reaction were then transferred to a series of tubes, each of which contained .1 vol of [y-3ZP]ATP and enough ATPyS or ADP to achieve the final concentrations indicated in Figure 7C. Samples for determrning P, release were withdrawn 1 min after addition of label. Assay for Open Complex Formation Open complex formation was quantitated by means transcription assay(Popham et al., 1989; Wedel et al., template was the supercoiled plasmid pJES528. This construction and characterization wrll be described tains a wild-type g/nA promoter regulatory region that

of a single-cycle 1990). The DNA plasmid, whose elsewhere, condirects syntheses

of a 155 nucleotide transcript containing no uridine residues. Omission of uridine triphosphate (UTP) during transcription allowed formation of the expected 155 nucleotide transcript and prevented synthesis of other transcripts. To assay inhibition of open complex formation by ATPyS or ADP, reactions were performed with components at the following final concentrations: 0.1 mglml acetylated BSA, 30 nM core RNA polymerase from E. coli, 50 nM as4 from S. typhimurium, 100 nM NTRCSIQF protein (constitutive), and 1 nM plasmid template. Components were added to buffer at 4% in the order indicated above. Mixtures were then warmed for 5 min at 37”C, and reactions were initiated by adding ATP, with or without ATPyS or ADP, to achieve a final ATP concentration of 1 mM in a total volume of 25 ~1. The final concentrations of ATPyS and ADP are indicated in Figure 7A. After 5 min, formation of open complexes was terminated by addition of heparin to a final concentration of 0.1 mglml. After an additional 5 min, guanosine triphosphate (GTP, final concentration 0.4 mM) and cytidine triphosphate (CTP, final concentration 0.1 mM; it contained 5 FCi of [@P]CTP [New England Nuclear]) were added to allow synthesis of transcripts. After 5 min, transcripts were precipitated, isolated by gel electrophoresis, and quantified as described. Radioactivity in transcript bands was converted to femtomole of transcript; the specific activity of transcripts was determined by multiplying the specific activity of CTP by64, the number of cytidine residues per transcript. To titrate ATP in the presence of a fixed concentration of ATPyS the following two modifications were made: formation of open complexes was initiated by adding a mix of ATP and ATPyS to achieve the concentrations indicated in Figure 78, and ATP was added to at least 0.1 mM (in addition to GTP and CTP) during synthesis of transcripts. Acknowledgments D. S. W. and J. B. contributed equally to this work and should be considered first authors. We thank Andy Wedel for assistance with transcriptionexperimentsandthephosphorimager, SuePorterfor help with the mobility shift assay, Anne Moon for some of the cloning, and Richard Burgess for the gift of core RNA polymerase. We are grateful to Mike Chamberlin, Caroline Kane, Jon Goldberg, Elizabeth Greene, and members of our laboratory for helpful discussions. J. B. was supported by a grant from lnstitut National de la Recherche Agronomique, France. This work was supported by Public Health Service grant GM38361 from the National Institutes of Health to S. K. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisemenl’ in accordance with 18 USC Section 1734 solely to indicate this fact. Received

June

20, 1991

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