A DNA architectural protein couples cellular physiology and DNA topology in Escherichia coli

A DNA architectural protein couples cellular physiologyand DNA topology in Escherichia coli

Robert Schneider, 1 Andrew Travers, 2

Tamara Kutateladze 3 and Georgi Muskhelishvili 1*1Institut fur Genetik und Mikrobiologie, LMU Munchen,Maria-Ward-Str. 1a, 80638 Munchen, Germany.2MRC Laboratory of Molecular Biology, Hills Road,Cambridge CB2 2QH, UK.3Institute for Molecular Biology and Biological Physics,Gotua str.16, 380060 Tbilisi, Georgia.

Summary

In Escherichia coli , the transcriptional activity ofmany promoters is strongly dependent on the nega-tive superhelical density of chromosomal DNA. This,in turn, varies with the growth phase, and is corre-lated with the overall activity of DNA gyrase, themajor topoisomerase involved in the elevation ofnegative superhelicity. The DNA architectural proteinFIS is a regulator of the metabolic reorganization ofthe cell during early exponential growth phase. Wehave previously shown that FIS modulates the super-helical density of plasmid DNA in vivo , and on bindingreshapes the supercoiled DNA in vitro . Here, we showthat, in addition, FIS represses the gyrA and gyrB pro-moters and reduces DNA gyrase activity. Our resultsindicate that FIS determines DNA topology both byregulation of topoisomerase activity and, as previouslyinferred, by directly reshaping DNA. We propose thatFIS is involved in coupling cellular physiology to thetopology of the bacterial chromosome.

Introduction

The topological fluctuations of DNA supercoiling inEscherichia coli differentially affect the expression ofmany genes and are believed to adjust cellular promoteractivity, both to the growth phase-dependent metabolictransitions and also to environmental changes (Prussand Drlica, 1989; Steck et al., 1993; Dorman, 1995; Tse-Dinh et al., 1997). One largely unexplored problem ishow the alterations in cellular physiology are coupledwith changes in the topology of the bacterial chromosome.

Besides the effects of cellular DNA transactions enga-ging processive DNA helicases, the total superhelical den-sity of eubacterial DNA is dependent on two importantfactors: the activity of the major topoisomerases, DNA gyr-ase and topoisomerase I, and constraint by DNA bindingarchitectural proteins (for review see Drlica, 1992). Theabundant DNA architectural protein FIS is synthesized inhigh amounts during early exponential phase of cellsgrown in rich medium (Ball et al., 1992; Ninnemann etal., 1992). FIS is thought to be involved in the adaptationof cellular metabolism to rapid growth by serving as anindicator molecule for the physiological state of the cell(Nilsson et al., 1992; Ninnemann et al., 1992). One impor-tant function of FIS is the activation of stable RNA (rRNAand tRNA) and related promoters on nutritional shift-up(Nilsson et al., 1990; Newlands et al., 1992; Champagneand Lapointe, 1998), with subsequent increase in thecapacity of the translational machinery. The stable RNApromoters also respond strongly to alterations in negativesupercoiling of DNA (Oostra et al., 1981; Lamond, 1985;Ohlsen and Gralla, 1992; Bowater et al., 1994; Free andDorman, 1994).

FIS regulates a large set of genes both by direct controlat the level of transcription initiation as well as by possibleindirect effects (Finkel and Johnson, 1992; Xu and John-son, 1995; Gonzalez-Gil et al., 1996). Besides specificallychanging the pattern of gene expression, FIS affects cellu-lar DNA transactions, including chromosomal replication,DNA inversion, phage integration/excision, DNA transpo-sition and illegitimate recombination (Gille et al., 1991;Filutowisz et al., 1992; Weinreich and Reznikoff, 1992;Betermier et al., 1993; van Drunnen et al., 1993; Cassleret al., 1995; Shanado et al., 1997). Many, if not all, ofthese DNA transactions are sensitive to the level of DNAsupercoiling (Mizuuchi and Nash, 1976; Mertens et al.,1984; Marians, 1987; Benjamin et al., 1996). Furthermore,even after the inactivation of specific FIS binding sites, inseveral of these systems the effect of FIS is retained, sug-gesting that FIS may act in a more global manner (Ball andJohnson, 1991; Betermier et al., 1993; van Drunen et al.,1993; Shanado et al., 1997).

By using different E. coli strains and plasmids with differ-ent replication control systems, we have shown that FISmodulates the dynamics of DNA supercoiling during thegrowth phase (Schneider et al., 1997). The growthphase-dependent topological fluctuations in DNA (Balke

Molecular Microbiology (1999) 34(5), 953–964

Q 1999 Blackwell Science Ltd

Received 5 July, 1999; revised 6 September, 1999; accepted 9 Sep-tember, 1999. *For correspondence. E-mail [email protected]; Tel. (þ49) 89 2180 6197; Fax (þ49) 89 21806160.

and Gralla, 1987) are strongly diminished in cells lackingthe fis gene. In particular, fis cells are impaired in their abil-ity to maintain a broad distribution of plasmid topoisomersthroughout the early to mid-exponential growth phase.The maintenance of this diversity of topoisomers requiresFIS binding and closely parallels the duration of fis expres-sion in the cell (Ninnemann et al., 1992; Schneider et al.,1997), suggesting that FIS constrains DNA and estab-lishes a distinct chromosomal architecture. It remainsunclear, however, whether the inferred reshaping ofDNA is solely responsible for the pleiotropic effect of FISon cellular DNA transactions.

Here, we show that FIS reduces the intracellular contentof DNA gyrase by repressing the promoters of gyrA andgyrB genes. This results in corresponding decreases inthe levels of GyrA and GyrB proteins and gyrase activity.Our findings indicate that, in addition to the effects of directbinding, the control of DNA topology by this architecturalprotein involves the modulation of the activity of one ofthe major cellular topoisomerases. More compellingly,

these results suggest that FIS links global DNA transac-tions with the physiological state of the cell.

Results

Effects of FIS on gyrA and gyrB promoter activity

In order to pursue the nature of the general effect of FIS onsupercoiling-responsive cellular DNA transactions, weinvestigated whether FIS affects the expression of thegyrA and gyrB genes, which code, respectively, for theA and B subunits of DNA gyrase (for review see Reeceand Maxwell, 1991). For this purpose, total cellular RNAwas isolated at intervals after the dilution of overnightcultures in fresh medium (this procedure is hereaftertermed nutritional shift-up) and the amount of the gyrAand gyrB transcripts measured in wild-type and fis cells.The measurements showed that, at the earliest testedtime point after the shift-up (5 min), the amount of bothmessages was lower in wild-type cells, the differencebeing more pronounced for gyrB (Fig. 1A–C). Afterwards,

Q 1999 Blackwell Science Ltd, Molecular Microbiology, 34, 953–964

Fig. 1. FIS represses gyrA and gyrB expression.A. Representative examples of Northern hybridization analysis of gyrA and gyrB mRNA respectively, are shown. eno RNA is shown as acontrol. The level of this RNA is independent of fis expression. Total RNA was isolated from CSH50 and CSH50Dfis cells at intervals after theshift-up as indicated.B. Quantitative Northern analysis of gyrA expression in CSH50 and CSH50Dfis cells. The data was derived from values obtained in sixindependent experiments.C. Quantitative Northern analysis of gyrB expression. The data was derived from values obtained in four independent experiments.

954 R. Schneider, A. Travers, T. Kutateladze and G. Muskhelishvili

the level of gyrA and gyrB messages increased in both celllines, but the relative abundance of both the gyrA and gyrBmessages in wild-type versus fis cells sharply decreasedafter 30 min, i.e. at the peak of FIS synthesis (Ninnemannet al., 1992). In mid-log phase (150 min after the shift-up),the amount of the gyrA and gyrB messages dropped butthe difference in their abundance between the wild-typeand fis cells remained significant.

We further investigated the in vivo transcriptionalresponse of the gyrA and gyrB promoters to FIS, usingthe plasmid-borne promoter–lacZ fusions, pGYRA1 andpGYRB1 (see Experimental procedures), and determinedthe b-galactosidase activity after their transformation intothe wild-type and fis cells. Consistent with the Northerndata, the fis cells showed significantly higher b-galactosi-dase activity than the wild-type with both constructs inexponentially growing cells, whereas the referencephage T7 A1 promoter cloned in the same plasmid back-ground (pA1lacZ) showed no significant differences in

activity (Table 1). Furthermore, the differences in thegyrA /B promoter activity between wild-type and fis cellswere not due to variations in plasmid copy number (seeExperimental procedures). Thus, the inactivation of fissimilarly affected two different variables – the amount ofchromosomal mRNA messages and the level of b-galacto-sidase protein expressed from plasmid-borne gyrA andgyrB promoters. The simplest assumption is that FISaffects the efficiency of transcription initiation at these pro-moters.

To test this assumption, we first carried out in vitro tran-scription reactions with linear gyrA and gyrB promoterfragments, using a multiple round run-off assay. We foundthat addition of FIS inhibited both gyrA and gyrB transcrip-tion in a concentration-dependent manner (Fig. 2A andB). Under the same experimental conditions, FIS did notinhibit the activity of the reference tyrT promoter (containingthe upstream activating region with FIS binding sites;Fig. 2C; Lazarus and Travers, 1993). Similar results wereobtained with supercoiled templates, but overall the tran-scriptional activity was much lower, as supercoiling by itselfimpairs the activity of the gyrA and gyrB promoters (data notshown; Menzel and Gellert, 1987).

Interactions between FIS and RNA polymerase atthe gyrA and gyrB promoters

We next investigated the gyrA and gyrB promoter regionsfor the presence of specific FIS binding sites, using DNaseI footprinting studies on linear promoter fragments in vitro.The addition of FIS protected the gyrA promoter DNA overan extended region between positions ¹16 and ¹68, clo-sely approaching the extended ¹10 hexamer element,and induced DNase I hypersensitivity around positions


Table 1. Expression of b-galactosidase in wild-type and fis cellscontaining gyrA /B promoter lacZ fusions.

ONa Exponential

CSH50 CSH50Dfis CSH50 CSH50Dfis

pGYRA1 9.000b 13.200 11.500 18.200(500) (500) (830) (400)

pGYRB1 27.700 39.200 38.200 72.300(1.100) (1.900) (3.100) (4.600)

pA1lacZ 81.350 82.500 92.500 94.500(2.800) (4.000) (2.400) (3.400)

a. ON,overnight cultures.b. Triplicate samples were assayed 2 h after the nutritional shift-up.Mean b-galactosidase activities are given in Miller units. Standarddeviations are indicated in parentheses.

Fig. 2. In vitro transcription from linear templates.A. Run-off transcription of the 208 bp gyrA promoter fragment (positions ¹136 to þ 72). The two messages initiating at the gyrA promoter areindicated. The concentration of linear gyrA template was 30 nM; the concentration of RNAP was 70 nM; the concentration of FIS (dimer) inlanes 2, 3 and 4 was 22 nM, 44 nM and 88 nM respectively.B. Run-off transcription of the 286 bp gyrB promoter fragment (positions ¹144 to þ 142). The concentration of linear gyrB template and FISwas as in (A), but the concentration of RNAP was 50 nM.C. Run-off transcription of the 310 bp tyrT promoter fragment (positions ¹150 to þ 160). The reaction conditions were the same as in (B).Note that at the highest used FIS concentration the stimulatory effect on transcription is lost, presumably because of FIS binding at the weaksite overlapping the ¹35 region of the tyrT promoter (Muskhelishvili et al., 1995).

FIS couples metabolism and DNA topology 955

¹16, ¹34 and ¹64 (Fig. 3A, lanes 5 and 6, and Fig. 3B).With increasing FIS concentrations, the extent of the pro-tected region was largely unaffected (compare lanes 5 and6). Such a protection pattern strongly suggested that spe-cific FIS binding in this region might inhibit transcriptioninitiation by steric hindrance. With the same DNA frag-ment, binding of RNA polymerase (RNAP) alone protecteda region between ¹19 and þ14 and, in addition, induced

several hypersensitive sites in the upstream region (at¹27, between positions ¹34 and ¹39 and around ¹62;Fig. 3A, lane 4), indicating that binding of polymerase dis-torts DNA in these regions. Addition of FIS displaced thepolymerase bound at the gyrA promoter, as judged bythe disappearance of both the polymerase-dependentprotection and DNase I hypersensitive sites (Fig. 3A,compare lanes 2, 3 and 4).

Specific FIS binding was also observed with the gyrBpromoter fragment. However, in contrast to gyrA, FIS pro-tected a region extending from ¹55 to ¹122 upstream ofthe core promoter, inducing DNase I hypersensitive sitesaround positions ¹70 and ¹104 (Fig. 4A, lanes 2 and3). Two sequences closely matching the FIS binding siteconsensus have been identified, centred at ¹62 and¹109 with respect to the start point of transcription(GNNYRNNA/TNNYRNNC; Lazarus and Travers, 1993;Fig. 4C). This location of FIS binding sites makes itunlikely that FIS directly interferes with polymerase bindingat the ¹10 and ¹35 elements of the gyrB promoter. Wealso footprinted the gyrB promoter fragment with RNAP,in the absence or presence of FIS. The results showedthat polymerase protection covered the start point andextended to near position ¹50, closely approaching theFIS binding site centred at ¹62 (Fig. 4A, compare lanes3 and 4). In addition, there was a polymerase-dependentalteration of the footprint around position ¹105 (lane 4,asterisk). As expected, occupation of the upstream regionby FIS did not hinder the binding of polymerase at the pro-moter (Fig. 4A, compare lanes 4, 5 and 6). However, inthe presence of FIS, the protection by polymerase wasnoticeably lessened at the positions ¹12 and ¹13, whereasthe protection in the region between positions ¹38 and ¹25was enhanced (compare lanes 4–6). Furthermore, anenhancement of protection was observed within the putativeFIS binding sites and in the intervening region betweenthem (compare lanes 2, 3–5, 6). Thus, although the simul-taneous binding of FIS and polymerase altered the inter-action of both proteins with the gyrB promoter DNA, thepolymerase remained stably bound to the promoter.These results argue strongly against the possibility thatrepression of gyrB transcription by FIS is a consequenceof a lowered affinity of the enzyme for the promoter.

We therefore investigated the effect of the addition ofFIS on the opening of the gyrB promoter under thesame conditions, using potassium permanganate as achemical probe. The addition of polymerase alone stronglyincreased the reactivity of thymine bases at positions þ3,þ2, ¹2, ¹6, ¹7 and ¹9 to potassium permanganate(Fig. 4A, lane 8). An additional weak hypersensitive sitewas observed near position ¹120. As potassium perman-ganate targets untwisted DNA, this result suggests thatbinding of polymerase leads to the untwisting of DNAwithin the region spanning the ¹10 hexamer and the


Fig. 3. Footprinting analysis of FIS binding at the gyrA promoter.A. DNase I footprinting of the gyrA promoter fragment. The regionsprotected by FIS and RNAP are indicated by black and grey bars,respectively. The concentration of FIS (dimer) was 15 nM in lanes 3and 6 and 44 nM in lanes 2 and 5. The concentration of RNAP was87.5 nM.B. Sequence of the gyrA promoter encompassing the FIS-protectedregion. The sequences matching the FIS binding site consensuswithin the protected region are underlined. The ¹10 element andthe start point of transcription are indicated.


start point of transcription. However, addition of FIShad no detectable effect on either the pattern or the extentof permanganate sensitivity caused by polymerase(Fig. 4C, compare lanes 8, 9 and 10). Thus, it is unlikelythat FIS reduces the efficiency of open complex formationat the gyrB promoter, but rather may affect a later step ininitiation.

To test this possibility, potassium permanganate foot-printing of the gyrB promoter fragment was carried outat a suboptimal temperature and in the presence of lownucleoside triphosphate concentrations to permit a slowisomerization of open complexes to initiation complexes.In this experiment, after preincubation of proteins with pro-moter DNA, the reactions were divided in two parts.Nucleoside triphosphates were added to one half of the

reactions for 5 s and then potassium permanganatewas added to all reactions for a further 10 s. Again, inthe absence of nucleoside triphosphates, the addition ofFIS had no effect on the reactivity of bases induced bypolymerase (Fig. 4B, compare lanes 1 and 3). However,addition of nucleoside triphosphates specifically increasedthe permanganate reactivity of thymines at þ 2, ¹7, ¹9and ¹10 with polymerase alone, whereas the presenceof FIS counteracted this effect (compare lanes 2 and 4).Taken together, these findings suggest that at the gyrBpromoter FIS prevents the isomerization of open com-plexes driven by substrate nucleoside triphosphates.

Thus, although the details of the inhibitory mechanismappear different, binding of FIS interferes with transcrip-tion initiation at both the gyrA and gyrB promoters. From


Fig. 4. Footprinting analysis of FIS binding atthe gyrB promoter.A. DNase I footprinting of the gyrB promoterfragment (left panel). The two regionscorresponding to sequences closely matchingthe FIS binding site consensus and the RNAPprotected region are indicated by verticalbars. Note that at higher concentrations FISalso weakly protects the region aroundposition ¹38 (lane 3). Potassiumpermanganate footprinting of the gyrBpromoter fragment (right panel). The reactivethymine bases (the bottom strand wasradioactively labelled) are indicated. Theconcentration of FIS was 44 nM in lanes 2, 5,9 and 11 and 88 nM in lanes 3, 6, 10 and 12respectively. The concentration of RNAP was250 nM.B. Potassium permanganate footprinting ofthe gyrB promoter fragment. The reactionswere carried out at 308C and nucleosidetriphosphates (NTPs) were added to the finalconcentration of 30 mM. The thymine baseswith increased reactivity to potassiumpermanganate on the addition of NTPs areindicated. The concentration of RNAP in lanes1–4 was 165 nM. The concentration of FIS inlanes 3 and 4 was 88 nM.C. Sequence of the gyrB promoterencompassing the FIS-protected region. Thetwo sequences closely matching the FISbinding site consensus are underlined. The¹10 element and the start point oftranscription are indicated.


these results, we infer that FIS acts as a transcriptionalrepressor of the gyrA and gyrB promoters.

Influence of the reinforcement of fis expression onthe intracellular level of gyrase

The repression of the gyrA /B promoters by FIS predictedthat the intracellular level of gyrase should be inverselycorrelated with the efficiency of fis expression. To testthis hypothesis we used two approaches. First, we carriedout multiple cycles of dilution of cells in fresh medium – aprocedure that supports a continuous expression of fis,such that high levels of FIS protein persist in the cell(Appleman et al., 1998). Using quantitative Western ana-lysis, we found that the repeated cycles of dilutionincreased the difference in the amount of GyrA and GyrBproteins (to roughly threefold for both GyrA and GyrBafter four to six dilution cycles versus 1.6-fold for GyrAand 1.77-fold for GyrB in a simple shift-up experiment)between the wild-type and fis cells (Fig. 5A). Thus, thecontinuous maintenance of high levels of FIS decreasesthe amount of GyrA and GyrB proteins. Under the sameconditions, the level of s70 protein showed no significantdifferences.

We next used cells lacking fis to test whether limitedoverexpression of fis from a plasmid would cause a reduc-tion in the levels of the GyrA and GyrB proteins. Theresults of these experiments showed that expression offis, but not of the DNA binding-deficient mutant fisR85C(Koch et al., 1991), significantly lowered the amount ofGyrA and GyrB proteins (2.9-fold and 2.4-fold respec-tively) in the cell (Fig. 5B). Taken together, these dataindicate that an inverse correlation exists between theexpression of fis and the content of DNA gyrase in the cell.

Determination of gyrase activity in cellular extracts

Finally, we asked how the observed difference in thelevels of gyrase between the wild-type and fis cells is

reflected in the supercoiling activity of corresponding cellu-lar extracts. Crude cellular extracts were prepared fromexponentially growing CSH50 and CSH50Dfis cells, aftermultiple dilution cycles as described above, and incubatedwith relaxed pUC19 plasmid DNA in the presence of ATP.The high-resolution agarose gel electrophoresis of plas-mid DNA demonstrated that the supercoiling activityof the extracts prepared from fis cells was significantlyincreased (Fig. 6). This result was reproducibly obtainedwith three different crude extracts, prepared both after sin-gle and multiple dilution cycles by using two different plas-mids (pUC19 and pBR322). Quantification of the amountof total extract protein required to achieve similar levelsof supercoiling indicates that, on average, the activity ofgyrase in fis cells is increased at least fourfold. We con-clude that the repression of gyrase production by FISleads to a decrease of intracellular supercoiling activity.


Fig. 5. Inverse correlation between the content of GyrA and GyrB proteins and fis expression.A. Comparative Western analysis of GyrA and GyrB protein synthesized in CSH50 and CSH50Dfis cells harvested after multiple dilutioncycles. The signals corresponding to GyrA and GyrB and the numbers of dilution cycles are indicated. The immunodecoration of s70 wascarried out with the same filter.B. Western analysis of GyrA and GyrB protein synthesized in CSH50Dfis cells after limited overexpression of fis (Schneider et al., 1997) fromplasmids containing wild-type fis or the DNA binding-deficient mutant fisR85C. Immunodecoration of the s70 protein was used as an internal control.

Fig. 6. The deletion of fis increases the DNA gyrase activity. Theprotein extracts were prepared from cells harvested after sixconsecutive dilution cycles and incubated with relaxed pUC19 DNA,as described in Experimental procedures. The strains from whichthe extracts were derived and the amount of total protein in thereactions are indicated. The samples were analysed on gelscontaining 0.3 mg ml¹1 chloroquine. The direction of migration isfrom top to bottom. Under these conditions the supercoiled DNAmigrates faster.


Discussion

The major conclusion of this study is that FIS repressesthe expression of both subunits of gyrase. This is sup-ported by several lines of in vivo and in vitro evidence.Our data show that, in vivo, fis expression correlateswith lower levels of both gyrA and gyrB mRNA and the cor-responding proteins, resulting in a reduction in intracellularsupercoiling activity. In vitro, FIS binding to the gyrA andgyrB promoters interferes directly with transcription initia-tion by RNAP indicating that, at these promoters, FIS actsas a transcriptional repressor.

Mechanism of gyrA and gyrB repression by FIS

The mechanism of the repressing effect of FIS on gyrAand on gyrB transcription differs. The gyrA promoterbelongs to the class of so-called ‘extended ¹10 pro-moters’, containing a 58-TGN-38 base sequence immedi-ately upstream of the ¹10 hexamer (Bown et al., 1997).At these promoters, the TG motif allows activity in theabsence of the ¹35 region, thus representing, in conjunc-tion with the ¹10 hexamer, the minimal sequence that canfunction as a promoter (Bown et al., 1997). Binding of FISprotects an extended region closely approaching and par-tially overlapping the ¹10 element of the extended gyrApromoter. Furthermore, the FIS-dependent DNase Ihypersensitive site at the position ¹16 targets the TGmotif of the extended ¹10 element and indicates that, inthis region, the DNA is distorted. In addition, in the upstreamregion, the protection by FIS overlaps two sequences thatshow RNAP-dependent DNase I hypersensitivity and thusare apparently involved in the RNAP–promoter interaction.All these observations strongly suggest that FIS repressesgyrA transcription by occluding the promoter.

There are few examples of repressors interfering withsteps in initiation subsequent to initial polymerase binding(see Choy and Adhya, 1996 and references therein). How-ever, these examples are significant for understanding thediversity of mechanisms that have evolved for negativecontrol. At the gyrB promoter, FIS interferes neither withpolymerase binding nor with the opening of the promoter.Instead, the mechanism of repression by FIS is probablycoupled to the intrinsic rate-limiting step of the gyrB pro-moter, which is proposed to be disabled at the stage ofpolymerase escape (Menzel and Gellert, 1987). Indeed,our potassium permanganate reactivity assays indicatethat open complexes form readily at the gyrB promoter.Moreover, although the alteration of the reactivity ofbases around the start point and within the ¹10 hexameron the addition of nucleoside triphosphates suggests anisomerization of open complexes, the signal does notextend downstream, indicating that there is little, if any,movement of the initiation bubble. Thus, the inhibitory effect

of FIS on the isomerization of open complexes may resultfrom an increase in the barrier of the rate-limiting step, mak-ing polymerase escape less feasible. The enhanced protec-tion of the gyrB promoter region in the presence of both FISand RNAP observed in our DNase I footprinting experi-ments is consistent with the notion of such a stalled ternarycomplex.

It is unclear at present how repression is effected bybinding of FIS at the upstream region of the gyrB pro-moter. One possibility, suggested by our DNase I andpotassium permanganate footprinting experiments, isthat polymerase makes contacts with the upstream regionof the gyrB promoter. If the architecture of the DNA micro-loop delimited by polymerase were to hinder polymeraseescape, then the stabilization of the microloop by FIS(Muskhelishvili and Travers, 1997; Muskhelishvili et al.,1997) could enhance this effect. An alternative but notmutually exclusive possibility is that direct protein contactsbetween DNA-bound FIS and RNAP lead to architecturalconstraints similar to those described for GalR–RNAPinteraction at the gal P1 promoter (Choy et al., 1997).

The general effect of FIS on DNA topology and DNAtransactions

The demonstrated coupling of fis expression and gyraseactivity indicates that DNA supercoiling directly respondsto a physiological effector signalling a metabolic reorganiz-ation. In our hands, the repressing effect of FIS on gyraselevels upon the shift-up is roughly twofold. However, bothan increase in the duration of fis expression by multiplecycles of cell dilution, as well as an overexpression of fis,lower the levels of gyrase further, suggesting that poten-tially the range of regulation is wider than suggested bya simple shift-up experiment. As the levels of FIS varygreatly, in response to both the nutritional quality of themedium and the growth phase (Ball et al., 1992; Nilssonet al., 1992; Ninnemann et al., 1992), we anticipate thatin the absence of any other influences, the levels of gyrasewill vary accordingly.

However, it is unlikely that the change in gyrase levelsdoes solely account for the difference in the overall super-helical density of DNA between the wild-type and fis cells.Our previous work demonstrated that FIS preferentiallybinds and stabilizes a subpopulation of moderately super-coiled DNA topoisomers during early and mid-exponentialphase (Schneider et al., 1997). Furthermore, in vitro, FISdirectly reshapes DNA, rendering it a poor substrate forgyrase. This direct binding effect may contribute to theobserved fourfold difference in supercoiling activity betweenthe wild-type and fis cell extracts. In line with such an effect,we have observed that binding of FIS constrains negativesupercoils in DNA (R. Lurz, R. Schneider, A. Travers andG. Muskhelishvili, manuscript in preparation).



Therefore, we suggest that by serving as an indicator forphysiological conditions, any fluctuations in FIS level willmodulate the topology of DNA, both by constraint and byaffecting the effective concentration of gyrase. More com-pellingly, FIS may play multiple roles in determining thearchitecture of prokaryotic chromatin. The proposed com-plex nature of the FIS effect is supported by studies on sev-eral supercoiling-responsive DNA transactions in which theinvolvement of specific FIS binding has been well documen-ted, but the effects could not be correlated with binding ofFIS at these sites (Ball and Johnson, 1991; Betermier etal., 1993; van Drunen et al., 1993; Shanado et al., 1997).The modulation of gyrase activity and DNA topology in thecell could provide an explanation for indirect effects of FISon supercoiling-responsive DNA transactions. Furthermore,the fact that many supercoiling-responsive promoters con-tain specific FIS binding sites (Lamond, 1985; Ohlsen andGralla, 1992; Free and Dorman, 1994; Sun and Fuchs,1994; Gonzalez-Gil et al., 1998) suggests that there aretwo variables that affect fis-dependent promoter activity:the superhelical density of DNA and the occupation of FISbinding sites (for detailed discussion see Travers and Musk-helishvili, 1998). Such a combination of direct and indirecteffects would allow for fine tuning of gene expression inresponse to changes in cellular physiology.

Implications for the homeostatic control of DNAsupercoiling

The level of unconstrained supercoiling in the bacterial cellis homeostatically controlled by the relaxing activity oftopoisomerase I and the supercoiling activity of DNA gyr-ase (Menzel and Gellert, 1983). In line with such a homeo-static control mechanism, we find that not only gyraseactivity is higher in crude extracts of fis cells, but the topoi-somerase I activity is also increased (R. Schneider and G.Muskhelishvili, unpublished observations). It is known thatan excessive supercoiling of DNA, owing to impairment ofcellular topoisomerase I activity, leads to compensatorymutations that reduce the supercoiling potential (DiNardoet al., 1982; Pruss et al., 1982). The negative superhelicaldensity of DNA is rapidly elevated on nutritional shift-up(Balke and Gralla, 1987). Thus, it is conceivable that animportant physiological function of FIS on nutritional shift-up is to counteract directly the generation of any excessivesuperhelical tension by gyrase that may drive inappropriateand potentially deleterious untwisting of DNA.

Experimental procedures

Chemicals and enzymes

Chemicals and enzymes used in this work were obtained fromcommercial sources.

Bacterial strains and plasmids

Bacterial strains used in this study were E. coli K12 deriva-tives. The genotype of CSH50 is ara D(lac pro) thi rpsL (Miller,1972). Construction of CSH50 Dfis is described elsewhere(Koch et al., 1988). This strain carries a kanamycin resistancegene substituted for fis. All strains were grown in 2× YTmedium (16 g of tryptone, 10 g of yeast extract and 5 g ofNaCl per litre, pH 7.4). Single colonies were isolated on YTplates (8 g of tryptone, 5 g of yeast extract, 5 g of NaCl and15 g of agar per litre).

For the construction of both the gyrA promoter–lacZ fusion,pGYRA1, and the gyrB promoter–lacZ fusion, pGYRB1, theconstruct ptyrTlac was used. ptyrTlac (containing the E. colityrT promoter–LacZ fusion; Helge Auner and G. Muskhelish-vili, unpublished data) was constructed by deletion of the bulkof the galK gene (838 out of 1194 bp) of ptyrTD150 (Lazarusand Travers, 1993), using restriction with Nar I and BstBI. ThelacZ gene derived from pTSV-11 (Wyckoff et al., 1986) wasthen ligated on a 3.4 kb SmaI–DraI fragment into Nar I/BstBI sites of ptyrTD150, yielding the plasmid ptyrTlac. As aresult of the cloning procedure, the first nine codons of thelacZ gene in ptyrTlac are substituted by 25 codons of galK.In the next step, the tyrT promoter of ptyrTlac was deletedby digestion with EcoRI/NruI. The gyrA promoter fragment,obtained by polymerase chain reaction (PCR) amplificationof the chromosomal gyrA promoter region from positions¹312 to þ37 with primers gyrA5311RI (58-CCAGAATTCCGCCGCTACAAC-38) and gyrA3RV (58-AGGTCGATATCCTAACCGCTATCCCTC-38), was cloned in the EcoRI/NruIsites of ptyrTlac, generating the gyrA promoter–lacZ fusionconstruct, pGYRA1. pGYRB1 was constructed by amplifica-tion of the chromosomal gyrB promoter region from positions¹355 to þ 30 with primers gyrB386RI (58-AATGAATTCCGCGATCGCCAGC-38) and gyrB1 (58-AGAATTTACGTGCAACGTTTCTCGCTC-38), and cloning the PCR-generated frag-ment in the EcoRI/SnaB1 sites of ptyrTlac. The SnaB1 sitein ptyrTlac is located < 100 bp upstream of the NruI siteused for pGYRA1 construction. The plasmid, pA1lacZ, wasconstructed by amplifying the phage T7 A1 promoter region(positions ¹65 to þ46) with primers A15RI (GGAATTCCAGATCCCGAAAATTTATCAAAAAG) and A13RV (CTGGATATCGACCCCGGTGTCGATTGG), and cloning the PCRproduct into the EcoRI/NruI sites of ptyrTlac.

Proteins

FIS was isolated as has been described previously (Koch andKahmann, 1986). E. coli RNA polymerase was purchasedfrom Pharmacia.

RNA isolation and Northern analysis

Samples of about 1010 cells were chilled on ice and collectedusing centrifugation. The RNA was isolated as described bySchmitt et al. (1990). The concentration of RNA in differentsamples was detected spectrophotometrically, normalizedfor each sample and verified on a reference gel, using rRNAas loading control. Either 15 or 30 mg of RNA per lane wereseparated on a 1% agarose gel in the presence of glyoxaland transferred to Hybond Nþ filters (Amersham). The filters



were stained with methylene blue in 0.3 M sodium acetate toverify that the transfer of RNA was quantitative. The filterswere hybridized overnight with a-32P-CTP-labelled probesmade by the Megaprime DNA labelling system (Amersham).An 898 bp gyrA gene fragment and a 620 bp gyrB gene frag-ment were used as template DNAs for labelling. The 898 bpgyrA and 620 bp gyrB fragments were generated usingPCR with the primers gyrA5 (58-GCGACCTTGCGAGAGAAATTACAC-38) and gyrA3 (58-CATACCGTCTTTGTCAGACTCGTC-38) and the primers gyrB5-1871 (58-CGTTGACGGAAGCTGACC-38) and gyrB3-2491 (58-GCCTGATAAGCGTAGCGC-38), respectively, using chromosomal DNA of E. colias a template. The eno probe was prepared using amplifica-tion of a 1283 bp internal fragment of the E. coli enolasegene, using the primers eno5 (58-AGTCTTATGCCTGGCCTTG-38) and eno3 (58-CATCGGTCGTGAAATCATCG-38).The signals were detected by phosphorimaging (PhosphorI-mager Storm 840, Molecular Dynamics) and quantifiedusing the IMAGEQUANT software, averaged and normalized tothe value obtained for the fis strain 5 min after the shift-up.

DNase I footprinting

The conditions of DNase I footprinting were essentially as hasbeen described previously (Muskhelishvili et al., 1995). ThegyrA promoter region (¹136 to þ72) was amplified by the pri-mers gyrAR5 (58-GTTTACCGAATTCTTTTCGGCATTC-38)and gyrAH3 (58-TCTTCCAAGCTTTTGACCGGTGTA-38).The gyrB promoter region (¹183 to þ30) was amplified bythe primers gyrB206RI (58-TGTGAATTCCTGATAGATGATTTTGCC-38) and gyrB1 (see above). For both PCR reac-tions, the E. coli chromosomal DNA was used as a template.The primers gyrAH3 and gyrB1 (bottom strands) wereuniquely end-labelled, using [g-32P]-ATP and T4 polynucleo-tide kinase. The fragments obtained were purified usingPAGE employing a neutral 0.5× TBE gel. The reactionswere assembled in a buffer containing 10 mM Tris-HCl,pH 7.9, 75 mM NaCl, 1 mM DTT, 10 mM ATP, FIS andRNAP as indicated, and equilibrated at 378 for 45 min beforeadding the mixture of DNase I and MgCl2 (to 2 mg ml¹1 and10 mM respectively) for 10 s. The samples were analysed ondenaturing 6% polyacrylamide gels and visualized using aphosphorimager. Protected and hypersensitive bands wereidentified using the Maxam–Gilbert G-ladder (Maxam and Gil-bert, 1977) of the same DNA fragments as a reference.

Potassium permanganate reactivity assay

The reactions for potassium permanganate reactivity assayswere assembled and processed similarly to those used forDNase I footprinting. After incubation at 378 for 45 min, 2 mlof 100 mM potassium permanganate solution was added for10 s to 20 ml of reaction mixtures containing DNA and pro-teins. The reactions were stopped by the addition of 2 ml of14 M b-mercaptoethanol, 8 mg of sonicated salmon spermDNA and sodium acetate to 0.3 M, precipitated with threevolumes of ice-cold ethanol and washed with 70% ethanol.The pellets were resuspended in 100 ml of 10% piperidineand incubated at 908 for 20 min. Then LiCl was added to0.5 M, the DNA precipitated with 3 volumes of ice-cold ethanoland washed at least twice with 100% and once with 70%

ethanol. The pellets were dried, dissolved in the loading dyeand analysed on 6% sequencing gels. The signals due to per-manganate reactivity of bases were visualized and quantifiedas described above.

Western analysis

For quantitative Western analysis, equal amounts of cellswere collected, resuspended in the sonification buffer(50 mM NaCl, 0.5% Triton ×100, 10 mM Tris HCl, pH 8) andcrushed by vortexing with glass beads at 48C for 20 min inthe presence of Complete (Boehringer Mannheim) proteinaseinhibitor. The amount of protein was determined using a BCAprotein assay (Pierce) and reference gels were run to inspecttotal protein load. Western blotting was carried out essentiallyas has been described previously (Muskhelishvili et al., 1995),except that gel electrophoresis was in 10% SDS–PAGE, andmethanol (10%) was included in the buffers during electro-transfer of proteins onto polyvinylidendifluoride membranes(Immobilon-P, Millipore). GyrA and GyrB proteins weredetected using GyrA- and GyrB-specific mouse antibodies(Lucent). s70 was detected using a RNAP s70 subunit-specificpolyclonal (goat) antibody (a generous gift of Hermann Heu-mann). Horseradish peroxidase-conjugated mouse (Pro-mega) and goat IgG-specific secondary antibodies wereused for detection with an ECLþPlus kit (Amersham). Theautoradiographs were scanned and quantified by NIH-IMAGE

software. The experiments were repeated at least threetimes to verify the obtained results.

In vitro transcription

Linear templates: multiple round run-off transcription reac-tions were performed with the linear 208 bp gyrA promoterfragment (see above), the 286 bp gyrB promoter fragment(positions ¹149 to þ 142) and the 310 bp tyrT promoter frag-ment (PCR amplified from position ¹150 to þ 160, usingptyrTlac as a template) in a buffer containing 10 mM Tris-HCl, pH 7.9, 10 mM MgCl2, 100 mM NaCl, 1 mM DTT, FISand RNAP as indicated, 0.5 mM each GTP, CTP and ATPand 0.05 mM [a-32P]-UTP in a 20 ml volume at 378C. The reac-tion was terminated after 2–5 min by the addition of an equalvolume of formaldehyde loading dye and heating. The sampleswere loaded on 6% denaturing polyacrylamide gels and ana-lysed using phosphorimaging as described above. The lengthof the transcripts was identified using corresponding sequenc-ing reactions as a reference.

Determination of the DNA supercoiling activity incellular extracts

Overnight cultures of CSH50 and CSH50fis::kan cells wereharvested after six consecutive dilution cycles in fresh dYTmedium. The dilution was 1:100 for the first cycle and 1:10for the following five cycles. The crude cellular extracts wereprepared according to DiNardo et al. (1982), except that Com-plete (Boehringer Mannheim) Proteinase Inhibitor was added.pUC19 DNA (250 ng), relaxed by vaccinia topoisomerase I(kindly provided by Karin Schnetz), was incubated with cel-lular extracts in the presence of 2 mM ATP and RNase (to



0.25 mg ml¹1). The concentration of total protein in cellularextracts was determined and normalized, using the BCA Pro-tein Assay Kit (Pierce). After incubation for 30 min at 308C, thereactions were stopped by the subsequent addition of EDTA(to 50 mM) and SDS (to 0.2%). Then proteinase K wasadded to 1 mg ml¹1 and incubation continued for 20 min at508C. After the addition of loading dye, the samples were sub-jected to high-resolution electrophoresis in 1% agarose gelscontaining 0.3 mg ml¹1 chloroquine, as has been describedpreviously (Schneider et al., 1997). The DNA in the gelswas stained with ethidium bromide and visualized under anUV light source.

Determination of plasmid copy numbers

The determinations of relative plasmid copy numbers in thewild-type and fis cells were carried out as follows. Total (chro-mosomal plus plasmid) DNA was isolated from wild-type andfis cells transformed with pGYRA1 or pGYRB1. The cellswere centrifuged and the pellets resuspended in a buffer con-taining 50 mM sodium phosphate and 50 mM NaCl, pH 8.0. Anequal volume of phenol was added, the suspension was vor-texed and incubated at ¹708 for 5 min. After centrifugationand a repeated round of phenol extraction, the total DNAwas precipitated by ethanol. Fifty nanogrammes of eachDNA was then transformed in DH5a cells, and equal aliquotswere plated onto solid media containing ampicillin to select forcells carrying the gyr promoter–lacZ fusions. The yield oftransformants per 50 ng of DNA isolated from wild-type andfis cells was 2420 versus 2300 for pGYRA1 and 2120 versus2210 for pGYRB1 respectively.

b-Galactosidase determinations

Overnight cultures were diluted 1:30 in fresh 2× YT medium.Samples taken at the indicated times were assayed for b-galactosidase activity, according to the protocol of Sadlerand Novick (1965). b-Galactosidase units were multiplied by1000 to make them equivalent to those of Miller (1972).

Acknowledgements

We thank Regine Kahmann for generous support, Rene Rostfor construction of pGYRA1 and Andrea Schultz for excellenttechnical assistance. This work was supported by theDeutsche Forschungsgemeinschaft through SFB 190.

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