Synergy between Escherichia coli CAP protein and RNA polymerase in the lac promoter open complex

J. Mol. Riol.

(1989) 206, 41--57

Synergy Between Escherichia coli CAP Protein and RNA Polymerase in the lac Promoter Open Complex David C. Straneyl-f-, Susan B. Straney’t and Donald M. Crothers1v2 ‘Department

of Molecular

Biophysics

and Biochemistry

2Department of Chemistry Yale University New Haven, CT 06511, U.S.A. (Received 8 February

1988, and in revised form 12 July

1988)

Characterization of ternary complexes containing an Escherichia coli lac promoter DNA fragment, CAP protein and RNA polymerase, separated on non-denaturing polyacrylamide of CAP against gels and footprinted in the gel slice, reveals a striking stabilization complex lacking RNA dissociation in the open complex, compared to the CAP-DNA polymerase. The stabilization is lost when half a helical turn of DNA is inserted between CAP and polymerase sites, but is partially restored with an 11 base-pair insert; stimulation of transcription parallels the stabilization effect. This behavior suggests a direct proteinprotein interaction. Comparison of initiation kinetics for wild-type and a mutant in which the P2 promoter has been inactivated shows that CAP both strengthens binding in the closed complex and accelerates isomerization to the open complex; the latter effect accounts for the bulk of the observed transcriptional activation.

competitor-resistant complex, RP,, with several base-pairs unwound or “open” (Kirkegaard et al., 1983; Siebenlist et al., 1980) flanking position 0 (relative to the start of transcription). Later it was found that upon addition of ribonucleotides, a stable initiated complex is formed, containing a 9 to 11 base RNA molecule, with concommitant loss of the polymerase subunit o (Carpousis & Gralla, 1985; Straney & Crothers, 1985). Complicating the picture is the finding that the Zac operon, like gal, has another transcriptional start site, P2, located about 20 bases upstream from the first site, Pl (Malan & McClure, 1984). Transcription from P2 is repressed by CAP, which binds in the -35 region of P2. In vitro, this is the major transcriptional start site in the absence of CAP, but no transcription from P2 can be detected in vivo (Spassky et al., 1984). The role of the P2 promoter in regulating the level of lac expression by Pl, along with the potential interaction between CAP and the P2 positioned polymerase, are unsettled issues (Malan & McClure, 1984; Spassky et al., 1984; Yu & Reznikoff, 1985; Gralla, 1985; Meiklejohn & Gralla, 1985). Malan & McClure (1984) proposed that CAP reduction of P2 occupancy indirectly stimulates the Pl promoter, while direct CAP stimulation of Pl has also been reported by Malan et al. (1984).

1. Introduction The mechanism by which CAP activates RNA polymerase has been postulated to involve either protein-protein interactions (Gilbert, 1976) or interaction through changes in DNA structure propagated between CAP and polymerase binding sites (Dickson et aZ., 1975; Wartel, 1977; McKay & Steitz, 1981; Ebright & Wong, 1981). Lack of definitive evidence linking either model to CAP activation hampers further efforts to elucidate the mechanism of activation or to account for the variation in distance between CAP and RNA polymerase sites in the various promoters which CAP activates (compared by de Crombrugghe et al. (1984)). In the basic model of transcription initiation (Chamberlin, 1974): Kb Ic, R + P=RP,=RP, k-2 RNA polymerase, R above, first binds to the promoter P, in a “closed”, competitor-sensitive complex (RP,), then isomerizes to an “open”,

t Present address: Department of Plant Pathology, Cornell University, Ithaca, NY 14853, U.S.A. 41 oo22~2836/X9/9,5cH)41-17 $03.00/O

0 1989 Academic PRRS Limited

42

I). (7. &raney

Another intriguing aspect of CAP stimulation is the unknown function of CAP-directed DNA bending, observed by decreased mobility on polyacrylamide gels (Wu & Crothers, 1984), electron microscopy (Gronenborn et al., 1984) and greater formation of ligatable minicircles (Kotlarz et al., 1986) of the CAP-DNA complex. The topology or energetics of the unusual interaction are expected to affect polymerase, but the functional role has not been elucidated. In the work presented here we utilize t,hc separation on native polyacrylamide gels of intermediat,es in wt#t lac transcription initiation which we reported earlier for the constitutive lac UV5

promoter (Straney & Crothers, 1985; S. Straney Rt Crothers, 1987). In addition, we describe a method for DNAase I footprinting of the complexes in the gel slice. We find strong stabilization of CAP binding

by

RNA

polymerase

in

the

Pl

open

complex. which depends on the helical phasing between the two sites, in parallel with CAP activation of transcription (seen initially by Mandecki & Caruthers, 1984) suggesting involvement of a protein-protein interaction between CAP and polymerase. In order to

eliminate

the

influence

of P2 in

transcription from Pl, we used a mutant promoter (i5) which has a weakened P2 promoter and a displaced CAP site due to the insertion of 5 bp between the - 10 and -35 region of P2. We measured the kinetic parameters for i5, and compared them t’o t,hose of the wild-type promoter when CAP is present’, in which case no binding of polymerasc to P2 is observed. We find that CAP increases the initial binding constant, R,, by a small amount, increases the forward isomerization rate constant, k,, by a larger factor, and has no effect on the rate of formation of initiated complex from the open complex.

2. Methods (a) Proteins

and DNA

The RNA polymerase holoenzyme was prepared as described by Straney & Crothers (1985). It was greater than 9OyA active as measured by open complex formation. CAP was prepared and kindly supplied by Dr Abraham Brown; it was purified using an affinity chromatography technique (Brown, 1987) and was approximately 70% active in specific DNA binding. The DNA fragment used was a 203 bp lar containing fragment, prepared from the plasmid described by Wu & Crothers (1984). (b) Formation

and electrophoresis protein-DNA complexes

of

RNA polymerase holoenzyme (5.5 x IO-’ M) and a 203 bp DNA fragment containing the lac promoter (2.4 x 10-s M) were mixed with or without preincubation with CAP (8 x 10d8 M) and CAMP (50 p(M). The reaction

t Abbreviations

used: wt. wild-type;

bp, base-pair(s).

et al. buffer was 40 mivr-Hepes (pH 8.0). 100 mM-potassium glutamate (pH %O), 10 miw-MgCl,, 0.1 mu-El)TA. 0.1 mnn-dithiothreitol, and bovine serum 0.5 mg albumin/ml. The reaction volume was normally 20 ~1. The reactions were incubated at 37°C for 10 min of CAP preincubation with the DNA, and 60 min with polymerase to allow complete formation of polymerase complexes. Competitors added t)o complexes prior to gel were either electrophoresis heparin (80 pg/ml). poly(dA-dT) (2 x lo-’ M in bp), or a 42 bp DNA fragment containing the CAP binding site (CAP site competitor). Gel electrophoresis was on a constant temperature gel apparatus maintained at 37”C, using a 4(p;, (w/v) polyacrylamide gel (l/40 bisacrylamide : ac~rplamide) wit’h (45 mM-Tris-borate (pH %3), 0.5 x TBE buffer 0.5 mM-EDTA) and 10 PM-CAMP recirculating in the buffer. The gel was run at 200 V until the xylem cyanol dye had migrated 10 cm. The DNA was either labeled \;ith [a-32P]dATP and Klenow fragment for visualization of gel complexes, or uniquely end-labeled for DNAasr 1 footprinting by cutting off t’he upstrea.m end with PvuTT. Initiated complexes were formed by incubating the preformed open complex reactions with 200 ,uM-(:pLh initiator, IO PM-ATP, IJTP and GTP, and 20 jlM-x1-()methyl CTP for 30 min at 37°C before loading onto the gel. (c) DNAase

I cleavage in, the gel slice

The gels were autoradiographed for 20 min at 25 “( : and the complex bands cut out in approximately 30 ~1 volume. Then 3 ,uI of a DNAase solution (containing: 10 mmTris . HCl (pH 8.0). DNAase I/ml. 0.4 Pg 2 mM-dithiothreitol. 50/O (v/v) glycerol, 0.5 mg bovine serum albumin/ml) was spread on the gel slice in an Eppendorf tube, which was then incubated for 45 min at 25°C. A solution of 50 mM-MgCl,, 50 InM-(:a(?i, (3 pi) was then similarly spread on the gel slice, which was then incubated at 37°C for 4 min before adding 15 ~1 of 0.1 M-EDTA to stop the DNAase I. followed in 4 min by SDS to 0.050/, (w/v), The gel slice was then electroeluted and the DNA precipitated with ethanol twice in the presence of 10 m&I-MgCl,. lyophylized from water. and dissolved in formamide loading dye. The samples were polya~~rylamitlt~ (I/20 bisacrylrun 011 IO?,(w/v) amide : acrylamide)/5O~,b (w/v) urea gels. The autoradiograph was scanned with a microdensitometer using the -89 position to normaliz? the scans. The complexes were made in 20 mM-potassium glutamate for Fig. 6 and 100 mmpotassium glutamate for those in Fig. 2. (d) Alul

cleavage to assay (‘A I’ ocrupa~q in open complex

The DNA fragment was labeled only on the upstream end so that cutting at the Alul site (-60) within the CAP site removes the label from the polymerasP-EDNA complex. This was achieved by labeling t.he Fnu4H (- 125)/RI (f67) fragment with [y-32P]ATP and kinase followed by cutting off the downstream-labeled end with HinfI ( + 55). CAP-polymerasr complexes were formed with this DNA in the st,andard conditions except onl? 20 mm-potassium glutamate was used to reduce the dissociation rate. AZuI (0.1 unit AluI/pI reaction) was added to the react,ion, followed after 30 s by the CAP site competitor DNA (42 bp CAP site-containing fragment, either 12 x lo-’ or 6x 10-’ M). Time point portions of the dissociation reaction were taken, added to reaction buffer with no potassium glutamate. 10O~~m-(~AMl (to

CAP

Protein-RNA

Polymerase

slow further CAP dissociation), and poly(dA-dT), and loaded onto the 4% polyscrylamide native gel described above. The amount of 32P retained in open or CAP complex was measured by cutting out the gel slices and counting them by Cerenkov counting. The data points were fitted to an exponential curve, using least-squares analysis. to calculate the tt of CAP dissociation. Although the competitor DNA was also cut by AU, the resulting half-sites bind CAP sufficiently to act as a competitor at these concentrations (Liu-Johnson et al., 1986).

43

interaction

terminated productive RNA, was cut out and measured by cerenkov counting. The exoII1 assay of open complex as formation was carried out on samples initiated described above without labeled UTP. ExoTII was added to the sample (1.6 units/p1 final concentration) incubated for 2 min, and the reaction was stopped and loaded as described above for exoII1. The time of exoII1 digestion, between 1 and 8 min, does not change the measured promoter occupancy. (h) Protein

(e) Excmuclease III

digestion

The DNA was uniquely labeled for exoTI1 digestion either from the downstream side of polymerase (used for measurement of Pl and P2 occupancies, DNA labeled as above for AluI cleavage), or from the upstream side (used for assaying CAP stability on 20 mmpotassium glutamate and CAP movement in 100 mM-potassium glutamate; RI/RI fragment labeled with [y-32P]ATP and kinase followed by cleavage with PvuII). In both cases, the unlabeled strand is uniquely protected from exoII1 digestion by filling in the ends with cr-thio nucleotides and phage T4 DNA polymerase before cutting with HinfI or PvuII to give the unique end label; we allowed incomplete digestion with these nucleases to leave some full-length DNA fragment that is not digested by exoII1 and so is used aq a standard for quantitating the amount of DNA loaded. Exonuclease III (16 units/pi) was added to preformed complexes (competed with poly(dA-dT)) at 1.6 units/p1 of reaction. Portions (4 ~1) were taken at the time points indicated and added to 16 ~1 of: 98% formamide, 15 mmNaOH, 12 mM-EDTA, xylene cyanol. Heated at 90°C and quenched on ice, these samples were run on 8 y0 polyacrylamide/50°/0 urea gels (l/20 bisacrylamide : acrylamide) until the xylene cyan01 had run 40 cm. The bands representing exoIII stops were cut out’ and measured by Cerenkov counting. The data points were fitted to an exponential curve using least-squares analysis, from which the rate of dissociation was derived and expressed as tt. (0 Construction

of phasing wLutants

The insertions between the CAP and polymerase sites were constructed by using the CAP-site cassette system constructed by Marc Gartenberg (Liu-Johnson et al., 1986; and unpublished results). Changes in the wt sequence at, - 4 1 and - 84 produce Sty1 sites into which synthesized fragments of the CAP site (42.mer) may be cloned. The synthetic DNA fragment used for cloning added either 5 (i5) or 11 (ill) bp of DNA between the -43 and -44 positions; the sequence of the insert was selected so as to mostly repeat the sequences between -45 and -49 in the wild-type. The construct was cloned using standard procedures. (R) Analysis

of CAP

stimulation

CAP st,imulation of Pl RNA synthesis was assayed after 30 min of CAP-DNA-polymerase preincubation. The same concentration of Pl-specific nucleotides was added as described above for the formation of initiated complex, but with 2OpCi of [32P]UTP and heparin (80 pg/ml final concentration) in a 20 ~1 reaction, and the reaction was allowed to transcribe for 30 min at 37°C. The reactions were stopped with an equal volume of 7 M-urea, 12 mM-EDTA, heated and loaded onto a 2036 polyacrylamide/50 y0 urea bisacryl(l/20 gel amide: acrylamide). The I I-mer RNA band, the

content of gel complexes

Open and initiated complexes were made as described in section (a), above except using g-fold higher concentrations of DNA, polymerase and CAP. After formation of the complexes at 37 “C, they were competed with 160 pg heparin/ml or 0.1 pg poly(dA-dT)/ml for 5 min, and then loaded into the non-denaturing gel. The resulting gel complexes were then cut out of the gel and subjected to electrophoresis on a 4% stacking/lo% separating denaturing SDS/polyacrylamide gel. as described by Straney & Crothers (1985). (if Kinetics

of open complex formation

Labeled i5 DNA (typically 5.6 x lo-” M) was incubated at 37°C for 2 min with no CAP or CAMP. Labeled wt DNA (typically 5.6 x IO-” M) was incubated for 10 min with 1.3 x lo-’ M-CAP and 50 PM-CAMP. Reactions were carried out in standard reaction buffer with 1 mg bovine serum albumin/ml. At time t=O, RNA polymerase was added, to give a final concentration between 1 x lOmE M and 2-5 x lo-’ M. At various times, 20.~1 samples were withdrawn and added to 3 ~1 of dye buffer (10x : 600% sucrose, with bromphenol blue and xylene cyanol) and 1 ~1 of 1.6 mg heparin/ml, to bind any polymerase that is free or in short-lived DNA complexes. The dye buffer and competitor were preincubated at 37°C. Samples were loaded immediately on a prerun, running, recirculating, 4% polyacrylamide gel as described above. Gel slices containing complexes were excised and counted in a scintillation counter, to determine the percentage of DNA in complex and free. (j) Equilibrium

measurements

i5 DNA (5.6 x lo-“M) was incubated at 37°C with varying amounts of RNA polymerase (final concentrations ranged from 2 X 1om9 M to 1 x lo-’ M) in the standard reaction buffer, for 4 or 5 h. as mentioned in Results. Dye buffer and heparin, as in open complex formation, were added, and the samples run on 4% gels and analyzed, as described above (see Results). (k) Kinetics

of open complex

dissociation

Labeled wt or i5 fragments (5.6 x 10-lo~) were incubated (with CAP and CAMP, 1.3 x lo-’ M and 50 FM, respectively. in the case of wt) with RNA polymerase (6 x 10V8 to 10 x lo-* M) at 37°C in the standard reaction buffer for 1.5 h. Then, at t=O, heparin (C,=Oa16 mg/ml) was added. Portions taken at various times were added to prewarmed dye buffer and loaded on a standard gel, and analyzed as described in Results and by S. Straney & Crothers (I 987). (1) fnitiated

complex formation

Labeled wt, or i5 fragments were incubated at 37°C as described above for open complex dissociation. At t=O, in

44

I). f’. Straney

addition to the heparin, a mixture of ribonucleotidrs. as described above, was added. Samples (20 ~1) were withdrawn at various times and added to prewarmed dye buffer and loaded immediately onto a standard gel. The

open and initiated

complexes had distinctly

different

mobilities, with the initiated complex (or complexes in t)hr case of i5) running faster than the open complex. Bands were analyzed as described in Results and by S. Straney & (kothrrs (1907).

3. Results (a) Protein-DNA complexes separated on polyacrylamide gels

Binding RNA polymerase to the wt lac promotercontaining 203 bp fragment and running the mixture on a native polyacrylamide gel produced bands of gel mobility similar to those we studied on the lac UV5 promoter (Straney & Crothers, 1985). As shown in Figure 1, RNA polymerase without (YAP formed one open complex (stable to competit>or DNA and therefore possessing a slow dissociation rate) and one closed complex (rapid dissociation rate with competitor) of greater gel mobility. Preincubation with CAP-CAMP, followed by incubation with polymerase, produced similar gel complexes. The closed complex with CAP had a decreased gel mobility compared to that without +CAP

+ Poi

et al CAP? presumably due to CAP bending of the I)NA, thus producing an anomolously slow mobility beyond t,he effect expected for the additional IO’!:, protein mass of CAP. Although t,he promot,er position (1’1 or P2) of closed complex is unknown, different positions occupied in the presenccl and absence of CAP would be unlikely to cause t’hr mobility shift, since PI and P2 open complexes run at the same gel mobility. In cont,rast, t,he open complex mobility did not differ with CAP addition, although CAP is bound in the open complex (from IINAase I footprinting and prot,ein gels of this complex: see below). The mobility of t)hc opcbn complex was similar to that of thr lac lrV5 0, open complex. Closed complex was observed on higher temperature gels than was the chasefor t’htb I’V5 promoter. for which closed complex was seen only on 5°C gels. The CAP complex. as shown in Figure 1, was stable to competition with poly(dA-dT), which is non-sprcilic for CAP, but was very sensitive t)o heparin, which dissociates (‘Al’ complex more rapidly t)han does cornpet,itor I)EA containing a (:AP sit’e of equal concentration, and does so within the 15 seconds of gel loading untfrr these conditions. Addition of rihonucleotides that allow PI -specific transcription of an O-methyl-C terminated 11-mer RKA (GpA dinucleotide initiator. specaific to the PI tCAP +Pol.

+rNTPs

Open -1nitloted Closed + Closed -

CAP

DNA I2345670

9

IO

II

Figure 1. Protein-DNA complexes using the wt promoter DNA fragment. llpon electrophoresis on a native 4%) polyacrylamide gel, the protein-DNA complexes produced from CAP-CAMP, RNA polymerase, or both are shown bound to a 32P-labeled 203 bp wt promoter DNA fragment with stated competitor treatment before gel loading. CAP ctAMP alone for 10 min incubation ( + heparin, lane 1; + poly(dA-dT), lane 2); polymerase alone for 5 min incubation (no competitor, lane 3; + poly(dA-dT), lane 4); polymerasc alone for 30 min incubation (+ poly(dA-dT), lane 5): CAPoAMP for 10 min then polymerase for 5 min incubation (no competitor, lane 6; + poly(dA-dT), lane 7; + heparin. lane 8); CAP-CAMP then polymerase for 30 min incubation (+poly(dA-dT), lane 9). Ribonucleotides added for an additional 30 min to allow initiation of 11-mer Pl RNA, +poly(dA-dT): CAP-CAMP then polymerase for 30 min, lane 10; polymerase alone for 30 min, lane 11. The 2 closed complexes are sensitive to the polymerase-specific competitor poly(dA-dT) due to their rapid dissociation rate, while the open and initiated complexes are not affected due to their slow dissociation rate. CAP complex is resistant to poly(dA-dT) due to the high specificity of CAP and the non-specific nature of this competitor DNA; however, CAP is rapidly dissociated from its DNA complex by heparin. The band below the open complex, polymerase alone, is a minor species with a dissociation rate intermediate between the values for open and closed complex.

CAP

Protein-RNA

Polymerase

Interaction

45

in Figure 2 as microdensitometer scans of the denaturing polyacrylamide gels. The CAP-DNA interaction on the top strand was seen as a series of protected bands (at’ minus 48, 50, 54, 58 and 63) and enhanced bands (at minus 45, 46, 55. 56, and 66), as seen initially by Schmitz (1981). The 10 bp phasing of enhanced bands suggests that the minor groove faces away from CAP at these sites (Spassky et al.. 1984), in agreement with molecular models for the complex. The open complex formed wit,h CAP displays a protection/enhancement pattern similar to a composite of t’he TJV5 open complex (lacking CAP), which contributes enhanced cleavage at - 23 and in the unwound region, together with the CAP site pattern seen in the binary CAP-DNA complex. These results imply retention of the (:A&DNA cont,acts in the ternary open complex. However, in contrast to the sensitivity of t,hr free (IAP-DNA complex t,o dissociation by heparin, the (‘AT’ footprint in the open complex is unchanged after 30 minutes of heparin competition, comparrd to the pattern seen when the complex is challenged with poly(dA-dT). Hence, polymerase-dependent interactions stabilize CAP binding in the open complex, slowing its observed dissociation from t.he CAP binding site. Fried & Crothers (1984b) determinrd that the dissociat,ion of CAP from its binding site is second-order with respect to DNA containing a competitor (:AP site, suggesting a direct attack of competit’or on the CAP-DNA c~omplrx. Our observed CAP dissociation rates WWC~ also dependent upon competitor DNA (see Table 1). W’e cannot, dissect t)he influences that slow t)he dissociation rat’? into fact’ors resulting from changes in competitor attack as distinguished from changes in simple CAP dissociation. Tt is likely that polymerase-CAP interactions that hold (‘Al) on the promoter would slow dissociation t)y both pathways. The initiated complex displayed a protection/enhancement pattern smaller than t,hat of the open complex, as seen previously in the U\‘5 promoter (Straney & Crothers, 1985), the protection between -44 and -9 being lost upon conversion t,o

start site, ATP, GTP, UTP and 3’OMe CTP; see Fig. 3) produced an initiated complex of increased gel mobility, containing the 1I-mer RNA but not the shorter abortive RNA products seen in solution. This was the same result as observed with the Pl promoter of Zac UV5 (Straney & Crothers, 1985). This initiated complex was only formed when CAP was present before open complex formation, as seen in Figure 1, indicating that the open complex without, CAP is incapable of Pl transcription despite the large yield of open complex without CAP. DNAase I and exonuclease III digestion studies discussed below show, as expected from previous studies (Malan & McClure, 1984; Spassky et al., 1984), that the majority of polymerases in this open complex were positioned at the P2 promoter and unable to transcribe the 1I-mer due to the specificity of the dinucleotide initiator and 3’.blocked CTP. The band below the open complex in the lanes containing polymerase alone has a dissociation t+ value of three minutes, and so is much less stable t,han thr open and more stable than the closed complrx. Tt s DNAase T footprint shows only two enhan& bands. at. + 15 and + 18 on the bottom strand and may represent another closed complex at PI. Alternately. this could represent a bound core polymerase; however, only purified holoenzyme was used and specific binding to the promoter is not expected without, the sigma subunit. (b) Il,VAasr

/ footprinting of the gel complexes: stabilization of CA P binding in the open complex

Because of our interest in assaying the presence and stability of CAP in the open complex following exposure to competitors including DNA and heparin, we developed a met’hod for DNase I footprinting in the gel slice so that competitor is separated from the complex before DNAase T treatment. (Heparin strongly inhibits DNAase T.) The protection/enhancement pattern of these complexes compared to bhose of bare DNA is presented

Table 1 (‘AI’

stimulation

ratio and CAP dissociation

Promotex

wt

8.7 0.82 2.6

i5 ill CAP--DNA complex

open complexes

CAP dissociation time from open complex tt (min)

(‘AP stimulation of Pl promoter (Ratio +CAP to -CAP) Open complex occupancy (ExoIII)

times in wt and mutant

RNA production (11 -mer) 8.3 0.83 1.6

AhIt digestion assa)

penetration of CAP site

4ooo ( 165) 1.6 19

I.7 0.059 0.22

6.7 (3%)

0.075

EXOIII

wt

t A/u1 digestion data in the presence of 6 x lo-’ M-CAP site competitor DNA. Data in parent,heses are at 12 x IO-’ M, the higher concentration producing faster dissociation and so a more accurate comparison of wt and CAP-DNA complex.

il.

C’. Stran.ey

et al.

-89 Reference

Bare

Figure 2. Scans of the DP;Aase I protection/enhancement pattern of the wt gel complexes. Shown are scans of denaturing polyacrylamide gels run with gel complexes treated with DNAase I. The regions of interaction are underlined, CAP being at the right and polymerase to the left. The positions numbered are those relative to the start site of transcription (+ 1). The top strand is shown. CAP enhances cleavage at positions -46. -56 and -66, protecting -54, and -61 and other sites between the enhanced bands. Polymerase in the open complex protects positions downstream from -45, enhancing cleavage at -22, as seen with Zuc IX5 and in the unwound region (- 1 to -7). Treat,ment of open complex with heparin for 5 min (shown) does not reduce the CAP protection, in contrast to the rapid dissociation of CAP from the CAP-DNA complex with heparin (complete in 30 s). The initiated complex shows a reduction in the upstream protection by polymerase, between -45 and -9. In contrast to the open complex. the CAP at the CAP site was removed by heparin treatment of the initiated complex.

initiated complex. The CAP site was still protected in initiated the complex challenged with poly(dA-dT), but as with the CAP-DNA complex, CAP dissociated rapidly from the initiated complex CAP site upon competition with heparin. The DNAase I pattern of the closed complex in the presence of CAP contained only the CAP site protection seen in the CAP-DNA complex, with no significant protection in the promoter region. Further experiments probing the closed complex

with exonuclease III digestion within the gel slice demonstrate at least two locations of bound polymerase: one producing an exo stop at approximately +34, possibly from the overlapping ~115 promoter (Peterson & Reznikoff, 1986), another at the Pl promoter, with an exo stop at +20. Therefore, the closed gel complex probably consists of at least two species of unstably bound polymerase, so neither site would be expected to produce a strong DNAase I protection due to

CAP Protein-RNA superposition of protected and unprotected in the mixture of species.

Polymerase Interaction

These constructs differ from those described by Mandecki & Caruthers (1984), who also inserted 5 and 11 base-pairs between the CAP and polymerase sites, by retention of the -75 to - 84 sequences and insertion further downstream in our mutants.

regions

(c) CAP-polymerase phasing mutants alter CAP stimulation and stabilization

(i) CAP stimulation of promoter activity in the phasing mutants

In order to study the nature of the interaction between CAP and RNA polymerase responsible for the stabilization of CAP in open complex, we constructed mutants with altered phasing between the CAP and RNA polymerase binding sites (Fig. 3). Five base-pairs are inserted in i5, rotating the face of the DNA interacting with CAP by half a helical t,urn relative to its position in wt; a whole helical turn is inserted in ill, which preserves the phasing of the two binding sites as in wt. These insertions were made at position -44, avoiding alteration of the sequences needed for CAP binding, positions -49 to -73 (Liu-Johnson et al., 1986), and those needed for RRiA polymerase binding in lac 1JV.5, determined to be -37 by resection experiments

ethylation

(Yu

& Reznikoff,

interference

1986) and

(Siebenlist

1980).

CAP site

-70

The ability

P-l

-35

-w

of CAP to stimulate

the Pl promoter

was measured both as increase in the yield of open complex, assayed by exoTI1 digestion, and by specific RNA production (quantitated in Table 1). Transcription was measured at Pl by formation of an 11-mer RNA specific to the Pl promoter due to the dinucleotide initiator used (GpA) and its size when terminated by 3’-O-methyl CTP. ExoIII digestion from the downstream side of the promoter halts at positions +20/21 in the Pl open complex and -4/5 in the P2 open complex (shown in Fig. 4), in agreement with the results of Yu & Reznikoff (1985). As quantitated in Table 1, the Pl promoter in the i5 mutant was repressed approximately 207/o by the addition of CAP. Experiments with a lac

- 39 by

et al.,

47

promoter

_, o

-40

+10

-35 P-2

CAP site

promoter

Polmerase -35 region

-.--em-‘----

-50

-30

-40

5’ CTCATTAGGCACCCCAGGCT-ITACAC

3’

TGGTA CGTCTAGGTAC

i 11

Figure 3. Construction of’ the phasing mutants i5 and ill. The upper panel shows the position of Pl and P2 promoters for the Zuc operon, with their corresponding - 10 and -35 control regions. The lower panel shows the sequences inserted between the CAP and Pl polymerase sites after position -44 in wt. Five base-pairs were inserted in i5, dephasing the binding sites by half a helical turn. Eleven base-pairs, close to a whole helical turn, inserted in ill retain the phasing between the binding sites. The base change at position -42 to a thymine was used to produce a Sty1 site in the cassette system of CAP site mutagenesis and so was present in the two mutants; this change does not alter CAP stimulation of Pl nor cause changes in the similarity to the -35 region consensus sequence. The CAP site is centered between -61 and -62; its boundary shown here is based on the work of Liu-Johnson et al. (1986). The -35 and - 10 regions are positioned by the Berg & von Hippel (1987) consensus sequence. The strong homology consensus is indicated by boxes (top panel) and continuous lines (bottom panel): the weak homology position is indicated by the broken line in the bottom sequence.

D. C’. Atraney et al

4x

-

-

wt,

I5 CAP

-

+

G

+ rNTPs +

+

-Initiated PI - Open PI -

P2 -

Figure 4. Polymerase occupancy at the Pl and P2 promoters assayed by exoII1 digestion from the downstream side of polymerase. Denaturing polyacrylamide gels separate the bands representing exonuclease III stops at the downstream edge of the polymerase open complex at the Pl (position +20/21) and the P2 (-4/5) promoters. The occupancy at each promoter is measured by the amount of DNA in each band relative to the total digestible DNA, normalized from the undigestible standard in each lane (top band). The addition of Pl-specific ribonucleotides (to make an 1 I-mer RNA) to wt open complex with GAP (+rNTPs lane) produces a downstream movement of polymerase in the initiated complex; in the absence of CAP, only the +20/21 Pl open complex band is chased into this initiated complex band while the -4/5 P2 band is not affected (data not shown). The G lane represents the substrate after the Maxam-Gilbert G cleavage reaction. construct in which the CAP site is replaced with non-specific DNA show a correlation of CAP repression of Pl transcription and CAP binding to the weaker CAP-2 site which overlaps the Pl promoter (Schmitz, 1981; Fried & Crothers, 1984a) as detected in 10 mlrr-Tris, 1 mM-EDTA polyacrylamide gel binding assays. We therefore ascribe the similar apparent repression of the i5 mutant to promoter occlusion by the CAP-2 site bound CAP

petitor

and conclude

(2) digestion

that

there

is no stimulation

In contrast to the lack of stimulation mutant

yields

partial that both assay methods.

compared

with

The insertion mutant

formation

stimulation measured for

by CAP.

in i5, the ill of Pl by CAP wild-type

by

base-pairs in the i5 P2 open complex in the absence of CAP (Fig. 4), and P2

severely

of additional decreased

run off transcription (lo-fold decrease from wt). The -35 region placement for P2 in i5 has greatly

reduced homology with the compiled consensus sequence of this region with two weak and one strong homology consensus positions (Berg & von Hippel, 1987), compared to four weak and two strong homology consensus positions in wt, and so is probably the cause of the decrease in activity. This fortuitous change allows study of the Pl promoter without interference from P2 open complex competition.

(ii) Measurement of CAP-polymerase mutant DNA

interaction on

The CAP-polymerase interaction which produces CAP stabilization in the open complex was measured in these phasing mutants in three assays: (1) DNAase I footprinting of the open complex at varying times after treatment with a CAP comDNA,

to determine

at a restriction

within the CAP site occupancy over a time

CAP-competitor tion past the combination

CAP

site

occupancy;

endonuclease

locus

to determine CAP site course of treatment with

DNA; (3) exonuclease III digesCAP bound at its site. The

of these three assays provides

informa-

tion on the positioning of CAP to nucleotide resolution (( 1) and (3)) while allowing quantitative measurement ((2) and (3)). DNAase I footprints of the open complexes of each mutant and wt are shown in Figure 5, with or without one to five minutes of competition with CAP site competitor DNA (the 203 bp lac wt promoter containing the CAP site). As shown above, the wt promoter retained the CAP-specific protection during competition for CAP. The ill mutant produced an open complex with the CAP site pattern unchanged from the CAP-DNA complex, but treatment with competitor DNA

_.... _.._......... -3

Il. CY.Straney

50

et al

CAP alone

(cl

Fig. 5. reduces the CAP-specific pattern over the five minutes competition. The i5 mutant binds CAP alone normally, but forming the open complex seems to have disordered the structure at the CAP site, as seen from the distorted DNA digestion pattern. This CAP site pattern was similarly abolished over 5 minutes of competition with CAP site DNA. The DNAase I pattern within the inserted region of i5 and ill, presumably at the CAP-polymerase boundary, changed upon addition of the RNA polymerase. The i5 insert contained enhanced bands at -46 and -48, which are present in the open complex regardless of CAP preincubation, and therefore are likely due to polymerase interaction at this site. The ill mutant had similar polymerase-induced changes in the DNAase I pattern in the insert, but also displays protection of position -52 in the insert, that was not present when either of the proteins was present alone. Ah1 digestion at the -60 position, within the CAP site, was used to quantitate CAP site occupancy over a time course of competition with CAP site DNA. Due to the single end-labeling of the fragment, digestion at this site was measured by loss of 32P counts from the open complex (quantitated in Fig. 6 and tt of fitted dissociation curves in Table 1). At CAP site competitor concentrations of 1.2 x 10e6 M, CAP in the wt open complex dissociates with a tt value 43-fold greater than the CAP complex alone. At half this competitor (a synthetic 42 bp CAP site DNA fragment) concentration, where dissociation is slower due to the second-order dependence of competitor on dissociation (Fried & Crothers, 19846), CAP dissociates from i5 open complex slightly faster than from CAP complex, whereas in the ill open complex it dissociates with a t+ value 2%fold greater than from CAP complex. CAP

dissociation from the wt open complex was much slower at this competitor concentration, producing a less accurate rate measurement. Using a wt promoter fragment with sequences upstream from the CAP site removed (HinPI cut at -81) does not significantly change the dissociation rate of CAP from wt open complex, eliminating the upstream sequences as a source of the stabilizing effect. Dissociation of CAP from the i5 open complex was not complete; the competitor-resistant portion of CAP in this open complex was further characterized in the exoII1 digestion assay described below. As a complement to the DNAase I and AluT dissociation assays, we measured the rate of exonuclease III digestion through position - 76, the upstream limits of the CAP-DNA interaction (Simpson, 1980), to assay stabilization of CAP. Passage through the - 76 position was measured by the disappearance of this band on denaturing polyacrylamide gels (Fig. 7, quantitated in Fig. 8 and Table 1). This was more rapid than CAP dissociation, suggesting a facilitation of dissociation by exoII1 processivity. Penetration through the CAP site in wt open complex was slowed 22-fold compared to CAP-DNA complex, while it was slowed three-fold through ill and was slightly accelerated through the i5 open complex. The i5 mutant displayed a minor exoII1 stop 5 bp downstream from the main CAP-specific stop, more stable to exoII1 penetration, only in the presence of polymerase and only in this mutant. This minor stop is consistent with repositioning a small portion (-4%) of CAP in the i5 open complex to a locus displaced from the CAP site, at the same distance from the polymerase as on the wt promoter, thus achieving the stabilization that we see for CAP in the wt open complex. This repositioned and stabilized CAP species of the i5 open complex would also explain the incomplete dissociation of

CAP Protein-RNA

Polymerase

51

Interaction

3007 240

250(“ii

&h x 9

2000

i I !

E 85 cn

150 -

I

V

\

3 IOO-

P i \

t 1 0

i

50 \ I 3

I 6

I 9

I 12

I 15

\

0-0

40

Time (min)

Time (min)

(b)

(a)

Figure 6. AM assay of CAP dissociation from open complex. (a) The amount of 32P label (counts/2 min) remaining in open complex is plotted against the time of AZuI nuclease digestion in the presence of 6 x 10m7 M-CAP site competitor. The cleavage at the AZuI site within the CAP site removes label from the subsequent gel complex and so measures CAP site occupancy. Dissociation curves shown are: wt promoter open complex (@), i5 open complex (V), ill open complex fragment) is (fl), CAP-DNA complex alone ( q ). The wt open cbmplex made with a shortened upstream end (H&PI/RI shown (0) with no associated curve. Curves are fitted with an exponential decay. (b) Similar AM digestion as (a) but in the presence of 12 x lo-’ M-CAP site competitor, which produces a more rapid CAP dissociation, and so allows more accurate measurement of the wt onen complex CAP dissociation rate (0) compared to that of the CAP-DNA complex (a).

CAP from this complex in the Ah1 assay and also the disordering of the CAP site DNAase I pattern of this complex. The relative dissociation values of CAP alone, and CAP with polymerase on wt and mutant promoters agree with those measured by the Ah1 assay described above, but are more rapid due to the more aggressive nature of the exoIIT probe. (d) CA P-polymerase interaction remains in initiated complex When open complex is allowed to transcribe 11 nucleotides to form a stably initiated complex, the upstream portion of the polymerase-promoter interaction is lost. This has been observed in lac UV5 (Carpousis & Gralla, 1985; D. Straney & Crothers, 1987), and is demonstrated here in the DNAase I footprint of Zac wt (Fig. 2). Since the upstream portion of polymerase is expected to be that which interacts with CAP, we have probed the initiated complex to determine if the CAPpolymerase interaction characteristic of open complex is retained with the polymerase movement, or if the interaction is lost. The protein content of initiated complex was determined after heparin treatment, which dissociates CAP-DNA complex

(Fig. 9). CAP protein was found to be present in the initiated complex; therefore CAP seems to remain associated with the polymerase complex in a dissociation-resistant manner, qualitatively similar to that seen with the open complex. This stabilized CAP is not positioned at the -61 CAP site since the DNAase I footprinting demonstrated no protection at this site in the presence of heparin. In order to determine if CAP is non-specifically bound to DNA when moved by initiated polymerase, we used exoII1 digestion to search for protein-DNA interactions in the initiated complex which might suggest such binding. Figure 10 illustrates exoII1 digestion from the upstream side of the promoter, with digestion pauses unique to open and initiated complexes. Open complex displays exo pauses at -76 (the CAP stop), -44, - 17 and - 12. Initiated complex contained exo pauses at -76, -24 and -6. Comparison of these results to lac UV5 without CAP shows the -24 initiated complex pause with CAP is unique, in that without CAP on UV5 there were no intermediate pauses between open complex ( - 44, - 34 and - 24) and initiated complex (-6) (D. Straney & Crothers, 1987). This intermediate -24 exo pause may represent CAP protein attached to the initiated polymerase

and

non-specifically

bound

to

DNA

52

Il. C!. Straney et al. wt

30

CAP

CAP

CAP POL.

DNA 45

30

i5

ill

CAP 15

15

30

-45

A’s

45

CAP

POL. 30

GAP IS

15

30

POL.

DNA 45

30

KS

600120

30

CAP 15

15

30

45

stop - 76

-43 -40

Figure 7. Exonuclease III digestion from the upstream side of polymerase: assay of penetration of the (‘Af’ complex. Denaturing gels show the exoII1 stop at the upstream edge of CAP in solution (-+) on the bottom strand; the position number of the stop ( -76 in wt) varies with the mutant due to the inserted DNA but is at, the same sequence in each case (-81 in i5 and -87 in ill). Early timepoints in the dissociation assay are shown for each mutant as labeled. The unique exoITT stop in the i5 open complex at base -76 is labeled ( > ). and only occurs in i5 with polymerasr present’. Bare DNA controls digested with exoII1 are shown in the lanes labeled DNA. ExoIII bands below the CAP stop represent pauses at the polymerase domains at -43. -40, -34. -18 and -17. similar wit,h or without CAP preineubation. The band at the top of the gel represents an undigestible DNA fragment used for normalizing the exoII1 bands in each lane; the band below this is the substrate band. here representing a fraction of the total which is protected by the presence of presumably end-bound polymerase. The A lanes represent the substrate after the Maxam Gilbert A > C cleavage rea,ction. The unlabeled lane in wt represents digestion of bare DNA.

of its normal binding site, It must be emphasized, however, that without comparison to a wt I’1 initiated complex lacking CAP (the yield of which is not great enough to enable the experiment) this band cannot be unequivocally assigned to the CAP protein. downstream

(e) i5 transcription

kinetics

In order to study PI transcription rates without CAP, the i5 mutant was used, thus eliminating the ” - 35 region” of the P2 site, while keeping the Pl site intact. Using our gel electrophoresis method to separate the open complex from free DNA, we measured the apparent on-rate of open complex formation, as described by S. Straney & Crothers (1987). Typical curves are shown in Figure 11. Plotting the apparent on-rate (l/z) versus the free

concentration of polymerase equilibrium and promoter (a quantity equal to the total amount of polymerase added, under our conditions of large polymerase excess) usually gives a curve such as the one shown for the wt promoter with CAP in Figure 12. However, for the i5 promoter, the apparent on-rate was the same for a wide range of (2 X 1o-8 M polymerase concentrations to 2 x lop7 M). We were unable to use lower concentrations of polymerase while maintaining the necessary excess of polymerase over promoter. We take the measured (constant) rate as an estimate for the isomerization rate lcz. In view of this difficulty, we measured instead the overall equilibrium constant of open complex formation, K= K&Jk-,. Since we had estimated lc,, and could measure k-,, determination of K allows us to find a value for Kb, the initial binding

CAP Protein-RNA

Polymerase

Interaction

53

Open c*ppo,.

+

Initiated -

+

+/-

Heparin

P’P 0.6

o-

a 5 0.5 0 ‘; c g 0.4 f al t ; 0

a

0.3

0.2

CAP

0.

( Time (mln)

Figure 8. Exonuclease assay of the stability of CAP in the open complex. The fraction of the total digestible 1)X,4 retained at the CAP-specific exoII1 stop ( -76 in wt, upstream boundary of CAP) is plotted against the time of exoII1 digestion. Digestions in the presence of RNA polymerase and CAP-VAMP are: wt promoter (O), prornot,rr (V), il I mutant promoter (0). i5 mutant Digestion in the presence of CAP-CAMP alone (0). The digestion time-course of the unique band in the i5 open complex (S). representing C,4P displaced 5 bp towards the polymerase, represents downstream approximately 4:,, of the open complex since only 50”/ of the DKA is in t,he open complex under these conditions in i5. Exponential curves are fitted with a least-squares program t,o obtain a t+ value.

constant. Increasing concentrations of polymerase were incubated with a fixed amount of DNA for four hours at 37”C, then half of each reaction mixture was loaded onto a standard complex gel. After

an additional

hour’s

incubation,

the second

half of each reaction was loaded onto another gel. Obtaining the same K values from reactions at the two incubation times assured us that we were at, or close to, equilibrium. Plotting (y’ open complex/% free DNA) tIersus the amount of free polymerase remaining (which equals the amout of polymerase added minus the amount of open complex) gave a simple linear plot whose slope was equal to K, the overall equilibrium binding constant of open complex formation. The first-order off-rate, k-,, was measured directly by sampling the amount of open complex remaining, at various times after adding heparin. Heparin, a polyanion competitor which sequesters free polymerase, prevents any dissociated polymerase from rebinding. The initial binding constant Kb was then determined by dividing the overall

Figure 9. Protein content of initiated complex after competition. Open and initiated complexes, made in the presence of CAP, were cut out of a non-denaturing gel then run on this denaturing SDS/polyacrylamide gel and to visualize the proteins. Mobility silver-stained standards of CAP (containing a minor contaminant of slower mobility) and RPU’A polymerase were run on the left. Competition with heparin for 5 min before running the complexes into the non-denaturing gel did not remove CAP from the open or initiated complexes, although similar treatment dissociates CAP-DN.4 complexes. As seen with the lac UV5 promoter (Straney & (:rothers, 1985). the initiated complex lacks the (r subunit. The band between cr and c( subunits is an artifact common to all lanes run from a treated polyacrylamide gel slice.

equilibrium constant of open complex formation by equilibrium constant. Wk- z> the isomerization Values for these quantities are given in Table 2. The first-order rate of initiated complex formation was measured as described by S. Straney & Crothers

(1987).

Unexpectedly,

two initiated

com-

plexes were formed. Roth contained the usual 11-mer of RNA starting at the Pl transcription start

site (results

not shown),

but were not further

characterized. The rate of formation of the sum of the two complexes is given in Table 2. (f) Kinetics

of wt with CAP

The kinetics of open complex formaGon and dissociation, as well as the rate of initiated complex formation on the wt promoter in the presence of CAP and CAMP were measured as described above for i5, except that the Kb value was determined by the usual dependence of apparent on-rate constant versus [R + P], as shown in Figure 12 (l/4 (S. Straney & Crothers, 1987). Figure II shows a typical curve and illustrates the overall enhancement of open complex formation at I’1 by CAP. The measured values for these constants are given

11. (‘. hkmeg

12

et al.

345678910

Twne (mm)

Figure 11. Kinetics of open complex formation. The fraction of free DNA versus time, for wt and CAP (@) or i5 (0). The concentration of polymerase in this reaction was 8 x lo-* M, the concentration of DNA (either wt or 5) was 5.6 x 10-l’ M. The concentration of CAP (used in the case of wt) was 1.3 x lo-’ M. Time refers to minutes after polymerase addition, [P]/[P] l=. is the fraction of promoter not in open complex at a particular time. Quantities were measured by the gel assay, as described in Results

0.64

Figure 10. ExoIII probing for CAP in initiated complex. ExoIII digestion from the upstream side of the promoter pauses at sites within the promoter. represented as bands on this denaturing polyacrylamide gel. The promoter positions of the exo pauses are noted on the side. In solution containing the open complex with CAP (lanes 3 to 6: digestion for 2, 6. 10 and 15 min, respectively), exoII1 pauses at positions: -76 (CAP stop), and -44, - 17 and - 12. In contrast, with solution containing initiated complex with CAP (lanes 7 to 10: digestion for 30, 45. 60 and 120s. respectively), exoII1 pauses at positions: -76 (CAP stop), -24 and -6. The formation of an exoII1 stop at -24, between those characteristic of open complex ( -44 to - 12) and initiated complex (-6), is not seen in the Zac UV5 promoter (I). Straney & Crothers, 1987); the -24 exoII1 pause is unique to the initiated wt promoter with CAP. This suggests that CAP has moved from the -76 exoII1 stop to the -24 stop as polymerase has entered the stably initiated state. Since these complexes were digested in solution, the remaining -76 CAP site exoII1 stop probably represents excess CAP complex remaining in solution. Lane 1 illustrates digestion with CAP alone over 5 min, showing complete penetration of the CAP site. Lane 2 is the Maxam-Gilbert A>C reaction as a sequence standard.

: 52 x 0.32 3

0.16

I

5

I

I

I

IO 15 20 [R+P] (xlO+Jw)

I

25

Figure 12. The apparent on-rate of open complex formation (l/t) as a function of the equilibrium concentrations of free DNA and RNA polymerase, [R + P]. The points on this plot were derived from curves such as the one shown in Fig. 11, for the case of wt and CAP.

CAP Protein-RNA

Polymerase Interaction

55

Table 2 Equilibrium

i5 wt +CAP

and kinetic parameters

KdM-‘)

M-‘1

4.4( kO.4) x lo6 1.q kO.2) x 10’

&6(f 1.5) x lo-5 7.6(+1.1)~10-~

in Table 2. It can be seen that the effect of CAP on Pl is mainly on the forward rate of isomerization, with a lesser effect on the initial binding.

4. Discussion In this study, we have observed a large stabilization of CAP binding due to the presence of RNA polymerase in the open complex. Viewed from the perspective of polymerase, the presence of CAP modestly stabilizes the closed complex and strongly accelerates isomerization to the open complex. Interactions in the open complex, presumably directly between polymerase and CAP, dramatically reduce the dissociation rate of CAP protein. Interaction between CAP and RNA polymerase has for some time been postulated to involve either direct protein-protein interactions (Gilbert, 1976), or DNA-mediated interactions in which binding of CAP alters DNA structure at the promoter and so facilitates polymerase binding or isomerization (Dickson et al., 1975; Wartel, 1977; McKay & Steitz, 1981; Ebright & Wong, 1981). Support has grown for the protein-protein interaction model, with the results of Kolb 8t But (1982) and Fried et al. (1983) showing no gross unwinding or H -+ Z transition in the CAP-DNA complex, along with the indications for co-operativity in interaction of CAP and polymerase with DNA demonstrated by Liu-Johnson (1986) and Ren et al. (1988). We sought to characterize the nature of the CAP-polymerase interaction in lac open complex by changing the phasing of the CAP and polymerase sites. In protein-protein interactions, the maintenance of binding sites on the same face of the DNA is essential, whereas DNA-mediated interactions would be expected to be primarily dependent upon the number of base-pairs between the sites. Our results demonstrate that the CAPpolymerase interaction in open complex is absent, and perhaps antagonistic, in the i5 mutant, in which the binding sites are out of phase by a half helical turn. However, the interaction is partially restored in the ill mutant, in which the distance between the binding sites is greater but the phasing is restored by a whole helical turn insertion. These results favor protein-protein interaction as the source of the observed CAP stabilization in wt open complex. Direct association of CAP and polymerase on the promoter reduces the dissociation rate of either protein from their binding sites: synergistic binding produces the observed stabilization of CAP binding without requiring inherent changes in the CAP-DNA interaction.

of transcription k-2(s-‘)

k,W’)

1.6(+0.2)x 1O-5 2+3( kO.3) x 1o-5

1.3(+0.6)x 1.3(+0.1)x

1O-3 10-A

An alternate model of DNA-mediated interaction, which would explain the phasing, is that the DNA upstream from CAP is bent acutely around CAP to interact with the back side of the polymerase in a way that produces CAP stabilization. This model is unlikely since removal of the DNA upstream of CAP (cutting at the HinPT site) does not reduce the CAP stabilization in the wt open complex in the Ah1 assay (Fig. 6). Our observation of partial CAP stabilization over the added 38 A (1 A =O.l nm) of 11 base-pairs requires either an unexpectedly long-range interaction, perhaps electrostatic, between CAP and polymerase. or that there is flexibility in one or more of the components. Such flexibility may derive either from elasticity of one or both of the proteins, or from the CAP-induced bending of the DNA which might physically draw together the two proteins. The latter model would offer a unique method for CAP to act over the various distances from the promoter as seen in different. systems; in fact. of the distances observed: most seem to differ by integral numbers of helical turns (41, 61 and 71 bp from start site to the center of CAP; de the partial Crombrugghe et al., 1984). Explaining activity of ill by flexibility in CAP-induced DNA bending would require that bending be increased relative to wt open complex to span the additional 38 A in ill. Experimental evidence for some DXA or protein flexibility in the CAP-polymerase interaction is seen in the Dn’Aase I footprints of the i5 and ill mutant, in which the enhanced cleavage at about -47 that results from polymerase binding appears in the inserted D?u’A where the CAP site pattern would begin in wt. This suggests either DNA bending in that region, or that the physical boundary of the polymerase can extend farther upstream than the downstream edge of the CAP complex in wt. The latter interpretation is consistent with ascribing t’he needed flexibility to the prot’eins, presumably polymerasr. (a) Comparison

of wt and 1’ T/5 gel complezes

The gel complexes described here for t#he wt Zac promoter are generally similar to those observed for transcription initiation on the lac UV5 promoter (Straney & Crothers, 1985), which include closed, open and initiated complexes. However. we do not observe two open complexes, as we had in UV5. Preincubation with CAP-CAMP changes the nature of the gel complexes in two ways. First, the position of the polymerase is mainly at the p2 promoter in

11. C’. Abaney

56

the absence of CAP, but at PI if preincubated with CAP, as expected from previous studies (Malan &Z McClure, 1984; Spassky et al., 1984; Lorimer $ Revzin, 1986). Second, the mobility of the closed complex is significantly reduced by the binding of CAP, as expected from the anomalously slow mobility resulting from CAP-induced DNA bending (Wu & Crothers, 1984), but in contrast t’o the similar mobility of open complexes with and without CAP. The degree of CAP-induced DNA bending in the open complex is under active investigation in our laboratory (S. Zinkel & D. M. Crothers, unpublished results). (b) Kinetic

effect of CAP

on PI

Using our assay, and mutant promoter i5, we were able to examine directly the effect of CAP on Pl transcription initiation. In the i5 mutant, all polymerase in open complex is bound at Pl, as is also the case on the wt promoter in the presence of sufficient amounts of CAP. As shown by DNAase T and exoTI1 protection (Figs 2 and 4), the interactions of RNA polymerase at PI in i5 appear, at this level, to be the same as in the wt DNA with CAP, in spite of the base change at -41 introduced for cloning convenience. We treat i5 as a “P2-less” wild-type in order to measure directly the kinet,ic effect of CAP on Pl. Unlike the kinetic measurements performed by Malan et al. (1984), we did not have to include any factors to take into account, promoter occupancy at P2. We find in contrast to their results that most of CAPS effect is on k,, the forward rate constant of isomerization (about a 14.fold stimulation), while a smaller effect (about 3*5-fold) is found on Kb. The total magnitude of the effect is 46-fold, which is similar to that seen in vivo (about 50-fold) by Beckwith et al. (1972). It should be acknowledged, however, that this agreement may be fortuitous, since, as we have stressed earlier (S. Straney & Crothers, 1987), the overall rates of transcription initiation in most in vitro systems are substantially lower than in vivo. Malan et al. (1984) have reported an 1l-fold increase in Kb, and no effect by CAP on k,. The reasons for this discrepancy are unclear. One explanation is that’ the complexes being measured may differ in the two assays. Those of Malan et al. (1984) are defined kinetically, by competence in abortive initiation, while ours are defined structurally, by their protein-DNA interactions (as measured by various footprinting techniques) and their stability to heparin. In addition, other evidence supports our finding that the major effect of CAP is on &. Garner & Revzin (1981) initially reported that without CAP, they could not obtain a poly(dA-dT)-resistant complex at the wild-type promoter, whereas they were able to do so with CAP. Since competitor resistance is a functional assay for the open complex, this suggested that it was at this step that CAP was acting. Furthermore, Lorimer & Revzin (1986). using exonuclease TIT as a probe, have

et al. shown that on the PI promoter wit,hout (‘Al’. polymerase will bind in a closed complex. but will not form an open complex without (:AP. This is in good agreement with our findings.

(c) Correlation of CAP interaction with CA 1’ stimulation

The changes we observe in the CAP-polymerase interaction in the phasing mutants correlate with the observed ability of CAP to stimulate transcription. These results agree with those of Mandecki & Caruthers (1984), who found similar CAP stimulation changes in vivo with 5 and 11 base inserts. A possible consequence of putative CAP-polymerase interaction in the open complex is an inhibition of polymerase movement from the promoter in subsequent transcription: either the CAP-polymerase or CAP-DNA contacts must be broken for polymerase translocation. We presented evidence suggesting that CAP does move with the polymerase during translocation to produce an 11-mer RNA. This implies a loss of CAP-DNA contacts while retaining the strong CAP-polymerase contact, since the initiated complex seems to maintain the CAP-polymerase interaction characteristic of open complex. The apparent relocalization of the upstream end of CAP to -24 suggests that CAP is still interacting with DNA in the initiated complex, but presumably in a nonspecific manner. CAP bound in this way does not produce a distinct DNAase I footprint in initiated complex. The energy required in breaking the CAPDNA bond for transcription to proceed does not seem to affect the measured rate of conversion from open to stable initiated but does increase the amount of abortive RNA produced before productive transcription begins (results not shown; LiuJohnson, 1986). The role of protein-protein contacts in bridging gene regulatory proteins is a very active field at present. Such contacts have been implicated involving ara (Dunn et al., 1984), 1 (Hochschild & Ptashne, 1986), lac (Deuschle et al., 1986) and gal repressors (Adhya & Majumdar, 1987). Bridging interactions also offer a possible model for eukaryotic enhancer elements working at a distance from the promoter (Ptashne, 1986). In these examples, protein-protein association provides cooperative effects at both sites, and loops the DNA interbetween binding sites. The protein-protein action which we inferred between CAP and polymerase spans a much shorter distance than in these examples, but analogy to the repressor systems can be seen in the helical phase dependence of CAP stabilization, and also in the potential role of CAP in enhancing DNA looping. Co-operative binding between RNA polymerase and regulatory proteins has been recently reported in other systems as well. Hwang & Gussin (1988) have measured a decreased amount of 2 repressor required for repression of an “up” mutation of A P,, promoter

CAP

and an increased amount of repressor “down” promoter mutant.

Protein-RNA

needed for a

We thank Marc Gartenberg for use of his cassette system of lac CAP site mutagenesis and CAPless mutant. This work was supported by a grant GM34205 from the National Institutes of Health.

References Adhya. S. & Majumdar, A. (1987). RNA Polym. Regul. Transcr. Proc. 16th Steenbock Symp. pp. 129-135, (Reznikoff, W. S., ed.), Elsevier, New York. Beckwith, J., Grodzicker, T. & Arditti, R. (1972). J. Mol. Biol. 69, 155-160. Berg, 0. G. & von Hippel, P. H. (1987). J. Mol. Biol. 193, 723-750. Brown, A. M. (1987). Ph.D. thesis. Yale University, New Haven, CT. Carpousis, A. J. & Gralla, J. D. (1985). J. Mol. Biol. 183, 165-177. Chamberlain, M. E. (1974). Annu. Rev. Biochem. 43, 721775. de Crombrugghe, B., Busby, S. & But, H. (1984). Science, 224, 831-838. Deuschle, U., Reiner, G. & Bujard, H. (1986). Proc. Nat. Acad. Sci., IJ.S.A. 83, 41344137. Dickson, R. C., Abelson, J., Barnes, W. M. & Reznikoff, W. S. (1975). Science, 187, 27-35. Dunn, T.? Hahn, S. & Schleif, R. (1984). Proc. Nat. Acad. Sci., U.S.A. 81, 5017-5020. Ebright, R. H. & Wong, J. R. (1981). Proc. Nat. Acud. Sci., IJ.S.A. 78, 40114015. Fried, M. G. t Crothers, D. M. (1984a). .J. Mol. Biol. 172, 241-262. Fried, M. G. & Crothers. D. M. (1984b). J. Mol. Biol. 172, 263-282. Fried, M. G., Wu, H.-M. & Crothers, D. M. (1983). Nucl. Acids Res. 11, 2479-2494. Garner, M. & Revzin, A. (1981). Nucl. Acids Res. 9, 30473059. Gilbert’, W. (1976). In RNA Polymerase pp. 193-206, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Gralla, J. D. (1985). Proc. Nat. Acad. Sci., (J.S.A. 82, 3078-308 1. Gronenborn, A. M., Nermut, M. V., Eason, P. & Clore, G. M. (1984). J. MOE. Biol. 179, 751-757.

Polymerase

Interaction

57

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