Capacity of ribosomal RNA promoters of E. coli to bind RNA polymerase

Cell, Vol. 10, 121-130, January 1977, Copyright 0 1977 by MIT

Capacity of Ribosomal RNA Promoters of E. coli to Bind RNA Polymerase

Karl Mueller, Christa Oebbecke, and Gisela Forster Zoologisches lnstitut der Westfalischen Wilhelms-Universitat Badestrasse 9 D-4400 Munster, Germany

Summary

The rate of in vitro transcription of the rRNA genes of E. coli is more than 20 fold higher than the averaged transcription rate of other genome segments of the same size. This “preferential transcription” of rRNA genes reflects a high efficiency of their promoters in chain initiation. We show that the high initiation rate at rRNA promoters results from a high rate of RNA polymerase binding to these promoters as measured by the formation of heparin-resistant RNA polymerase-DNA complexes. The results indicate that the preferential binding of RNA polymerase to rRNA promoters is mainly due to their large binding capacity rather than to a high rate constant of polymerase binding to a single binding site.

The polymerase binding capacity of rRNA promoters was estimated from the number of rRNA chains initiated by heparin-resistant complexes under conditions of template saturation and from the number of rRNA transcription units participating in the binding reaction. At least 30 RNA polymerase molecules were found to be protected from heparin per rRNA transcription unit. The rest of the genome (99.4%; possibly sufficient to en- code 4000 nonribosomal RNA species) protects under these conditions 2000 enzyme molecules. These results suggest that a high multiplicity of RNA polymerase binding may be responsible for the high efficiency of rRNA promoters. The validity of this hypothesis is discussed.

Introduction

Initiation of transcription is controlled by the interaction of promoter regions of the DNA with the enzyme RNA polymerase (E.C. 2.7.7.6.). A promoter region could simply be conceived as a specific sequence of approximately 40 base pairs (bp) which is recognized by RNA polymerase as a binding site and includes the starting point of transcription. This concept would agree with the fact that DNA sequences protected from DNAase by binding to RNA polymerase contain specific starting sites of transcription (Heyden, Nusslein, and Schaller, 1975; Pribnow, 1975). More complex promoter models, however, have been constructed (Schafer,

Zillig, and Zechel, 1973a; Dickson et al., 1975) to account for certain features of promoter function, particularly for the phenomenon of multiple binding of RNA polymerase to promoter regions: measurements of the stoichiometry of complex formation between RNA polymerase and bacteriophage genomes (Mueller, 1971; Schafer et al., 1973b) or individual promoters of phage lambda (Willmund and Kneser, 1973) indicated that a single promoter can bind several RNA polymerase molecules in form of heparin-resistant or poly(dAT)-resistant complexes. The multiplicity of binding was found to be characteristic for the type of promoter (Schafer et al., 1973b; Willmund and Kneser, 1973) and, in the case of the gal promoter of phage hpgal, to depend upon the presence of catabolite gene- activating (CGA) protein and cyclic AMP (Willmund and Kneser, 1973). Different promoters thus appear to contain composite binding regions of different- and in some cases, perhaps regulated-binding capacities.

We see the significance of variable binding capacities of promoters in the possibility that they may be responsible for variations of promoter efficiency in vivo, where the rate of chain initiation is limited by the concentration of free, active RNA polymerase (Bremer and Dalbow, 1975). According to this concept, it would be expected that highly efficient promoters such as those of ribosomal RNA genes of bacteria (30 chain initiations per min at steady state; Bremer and Dalbow, 1975) have a higher binding capacity than those of genes coding for mRNA (for example, derepressed trp operon at steady state: 1.1 initiations per min; Baker and Yanofsky, 1972). Here we report measurements of the binding capacity of rRNA promoters of E. coli under in vitro conditions in comparison with measurements of the binding capacity of the total E. coli genome. The results are consistent with the concept described. They show that the high efficiency of rRNA promoters in vitro is paralleled by a large capacity for RNA polymerase binding. After completion of this work, qualitatively very similar conclusions were reported by Venetianer, Sumegi, and Udvardy (1976).

Results

Experimental Concept Formation of RNA polymerase complexes with rRNA promoters was experimentally defined by the resistance of template-bound RNA polymerase against heparin and its ability to transcribe rRNA genes. To measure the binding capacity of rRNA promoters of the E. coli genome, the conditions of the following procedure had to be established:

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-Complexes of RNA polymerase and E. coli DNA are formed under conditions of template saturation with enzyme. -The excess of free RNA polymerase is selectively inactivated by heparin (Zillig et al., 1970; Schafer et al., 1973a). Eventual instability of the enzyme DNA complexes has to be considered. -After treatment with heparin, the mixtures are incubated with substrates for RNA synthesis. Each heparin-resistant RNA polymerase-DNA complex may thus initiate one RNA chain, because reinitia- tion after chain termination is prevented by heparin (Zillig et al., 1970; Schafer et al., 1973a). -The maximum number of enzyme molecules protected from heparin by binding to a rRNA promoter is determined from the number of rRNA molecules initiated in the presence of heparin and from the number of rRNA transcription units participating in the binding reaction.

Resistance of RNA Polymerase-rDNA Complexes against Heparin RNA polymerase is completely inactivated if it is added to reaction mixtures containing E. coli DNA and heparin at a concentration of 0.2 mg/ml each (data not shown). It can, however, become resistant against heparin by binding to E. coli DNA. This is shown by the experiment of Figure 1, where RNA

1.0 I I

x %

x E ,, ------X-- j

c I ----x --_______

-------7 P 8 g 05.. 0:

t? *-Otg

3

O1,,_._., Hepar~n concentration ( mg/ml )

Figure 1. Resistance of RNA Polymerase-DNA Complexes against Heparin

Binding mixtures were prepared, each containing 16.6 Fg of RNA polymerase and 9 rg of E. coli DNA in binding buffer (final volume 220 pi). After IO min of incubation at 37”C, different amounts of heparin were added to the mixtures in a volume of 10 ~1. Tran- scription was started 2 min later and was stopped after a 10 min period of RNA synthesis by treatment with phenol. Total incorporation of 3H-UMP into RNA was determined from the amounts of acid-precipitable radioactivity in the aqueous phases after phenolization. Incorporation of 3H-UMP into rFtNA was measured in the presence of internal standard rRNA (0.16 pg of 14C-rRNA per filter, 27670 cpm/pg). The fraction of rRNA and JH-UMP incorporation into total RNA are plotted against the concentration of heparin in the binding mixture. (O-O) incorporation of 3H- UMP into total RNA/reaction mixture; (x---x) fraction of rRNA (% total RNA products).

polymerase was incubated with E. coli DNA before heparin was added in varying concentrations. It is seen that the presence of heparin in the reaction mixtures reduces the rate of total RNA synthesis by preventing consecutive chain initiations, and that there is a well defined fraction of activity which is almost insensitive to the inhibitor. It represents the activity of RNA polymerase-DNA complexes formed during the period of preincubation.

Some conclusions concerning the function of rRNA promoters can be drawn from the results of Figure 1: approximately 15% of the RNA products are transcribed from rDNA, independent/y of the presence of heparin. If the rate of rRNA synthesis is related to the amount of rDNA which constitutes only 0.66% of the total template (see last section of Results), a “preference” of RNA polymerase for the transcription of rRNA genes becomes evident in the sense that these genes support a >20 fold higher rate of transcription than average template sections of the same size. RNA chain length measurements presented below (Figure 7) exclude the possibility that this phenomenon is due to differences in the rates of elongation or termination of ribosomal and nonribosomal RNA chains. It can be stated, therefore, that rRNA promoters are highly efficient in chain initiation. More precisely, their efficiency is due to a high rate of RNA polymerase binding rather than a high intrinsic rate of chain initiation, since preferential transcription of rRNA genes occurs in reaction mixtures which received heparin.

To test the stability of RNA polymerase-DNA complexes, they were exposed to heparin for different periods of time before transcription was started (Figure 2; corrections for hybridization efficiency were omitted for better judgment of significance). The activity of RNA polymerase in the complexes is seen to remain constant during a 30 min period of incubation with heparin. The complexes are obviously homogeneous in their stability; those formed with rRNA promoters are as stable as those formed at other binding sites. The slight increase in activity during the first minutes of incubation with heparin was reproducible. It may be due to conver- sion of “delayed starters” to “immediate starters” (Schafer et al., 1973b). In standard binding tests, the substrates were therefore added to the binding mixtures 2 min after mixing with heparin.

Kinetics of rRNA Synthesis by RNA Polymerase- DNA Complexes There is no direct evidence to answer the question how long polymerase-DNA complexes should be incubated with substrates to assure that chain initiation at the rRNA promoters is completed. How- ever, some indications may be derived from the

RNA Polymerase Binding Capacity of rRNA Promoters 123

B K

Ob 5 30 T~rlnOe Ofte~50dd~t~02n00f hepZdrn hn)

Figure 2. Stability of RNA Polymerase-DNA Complexes

Standard binding mixtures were incubated for 10 min at 37°C. Heparin was then added to each mixture (50 pg in a volume of 10 PI), and the incubation was continued. At different times after heparin addition, transcription was started by addition of substrates. After a 10 min period of RNA synthesis, the mixtures were treated with phenol, and samples (200 ~1) of the aqueous phases were hybridized with DNA on filters which had been preincubated in the presence or absence of nonradioactive rRNA. The symbols represent average values of duplicate determinations. (A) (O-O) 3H radioactivity of RNA hybridized in the absence of competitor rRNA; (B) (O-O) 3H radioactivity of nonribosomal RNA hybrids (that is, of RNA hybridized to “blocked” DNA); (C) (A-A) 3H radioactivity of hybridized ribosomal RNA, calculated as the difference of the values of (A) and (8).

kinetics of r-RNA synthesis shown in Figure 3. While the rate of nonribosomal RNA synthesis declines steadily from the beginning of the reaction, rRNA synthesis proceeds initially at an increasing rate. This effect suggests that delayed chain initiations occur at the rRNA promoters throughout the first 5- IO min of the reaction. As a consequence of late initiations of rRNA chains, the fraction of rRNA products (broken line in Figure 3) increases initially up to a reaction time of approximately 10 min. Thus if we require every RNA polymerase molecule bound to a rRNA promoter to initiate transcription, the period of RNA synthesis following complex formation should last at least 10 min.

It has been noted previously by Pettijohn (1972) and Travers, Baillie, and Pedersen (1973) that the rate of rRNA synthesis gradually increases during the first 10 min of the reaction started by addition of enzyme. As an explanation of this lag of transcriptional activity, Travers et al. (1973) suggested that the structure of rRNA promoters requires approximately 10 min of incubation at 37°C to be fully “opened.” This hypothesis cannot explain the kinetics of Figure 3, since in this experiment, RNA synthesis was started after preincubation of the enzyme-template mixture for 10 min at 37°C. The results of Figure 3 indicate that initiation of rRNA chains is limited by a relatively slow reaction which

Time of RNA syntheslsimml

Figure 3. Kinetics of rRNA Synthesis by RNA Polymerase-DNA Complexes

RNA polymerase-DNA complexes were formed in standard binding reactions. RNA synthesis was then started by adding to each mixture 300 +I of a 3H-UTP substrate solution containing 50 pg Of

heparin. Incubation at 37°C was continued. The reactions were terminated after different transcription periods by treatment with phenol. The contents of total JH-RNA and W-rRNA were measured. They are expressed in the figure in terms of ‘H radioactivity incorporated per reaction mixture (averages of duplicate determinations). (A-A) radioactivity in rRNA; (O-O) radioactivity incorporated into nonribosomal RNA (= total acid-precipitable RNA minus rRNA); (x---x) fraction of rRNA (% total radioactivity incorporated).

follows the rapid formation of heparin-resistant complexes (see below, Figure 4).

Conditions for Saturation of rRNA Promoters with RNA Polymerase The extent of RNA polymerase binding to DNA de- pends mainly upon two variables: the duration of the binding period and the weight ratio of enzyme/ DNA. In the following experiments, the effects of these variables on complex formation were studied to select conditions for saturation of the binding sites with RNA polymerase.

Figure 4 shows the kinetics of binding at two different weight ratios of RNA polymerase/DNA. At a weight ratio of 2 (corresponding to approximately 350 bp per enzyme molecule), almost all heparin- resistant complexes are formed within 2 min after addition of RNA polymerase to the template (Figure 4, curves A and B). At a IO fold reduced DNA concentration (weight ratio of 20; definite excess of RNA polymerase), biphasic kinetics of complex formation are observed (Figure 4, curves C and D): after a period of rapid polymerase binding, there is a period of relatively slow complex formation with <l/20 of the rate of rapid binding. We have not investigated which level of complex formation would finally be attained by the slow mode of binding, because it contributes very little to the complexes formed at the lower weight ratios at which template saturation is reached (see below). AC-

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Figure 4. Kinetics of Formation of Heparin-Resistant RNA Polym- erase-DNA Complexes

RNA polymerase (16.8 pg) was added to binding mixtures (37°C) containing either 8.5 rg of E. coli DNA (curves A and B) or 0.85 pg of E. coli DNA (curves C and D). Heparin was added (50 pg per mixture in IO ~1) at different times after the start of complex formation, and transcription was started 2 min later by addition of substrates. After a 10 min period of RNA synthesis at 37”C, the reactions were stopped by phenolization. The total acid-precipitable RNA products and the amounts of rRNA and nonribosomal RNA were determined. (A and B) Results of reactions with 8.5 fig of DNA. (A) ( 0- 0) 3H radioactivity in nonribosomal RNA; (8) (A- --A) 3H radioactivity in rRNA. (C and D) Results of reactions with 0.85 rg of DNA. (C) (O-O) 3H radioactivity in nonribosomal RNA; (D) (A- - -A) 3H radioactivity in rRNA.

cording to these results, the choice of a 10 min period of complex formation for binding tests would ensure that rapid complex formation is completed.

To determine the saturating weight ratio of RNA polymerase/DNA, two sets of binding mixtures with constant concentrations of RNA polymerase (8.5 pg per mixture or 16.8 pg per mixture, respectively) were incubated with variable amounts of E. coli DNA, and the activities of the heparin-resistant complexes were measured. It is seen in Figure 5 that in the range of limiting template concentrations, the rates of RNA synthesis rise with increasing DNA concentration up to a maximum. This maximum indicates the upper limit of template saturation. It is proportional to the enzyme concentration (compare the position of the peaks in Figure 5) and corresponds in both sets of binding mixtures to a weight ratio (enzyme/DNA) of 3 to 4. At DNA concentrations above the limit of saturation, the rate of RNA synthesis does not remain constant but decreases. This inhibitory effect of high DNA concentrations is not understood. Experiments with

“ _ ~ ~

0 5 io i5

Amount of DNA (pgl

Figure 5. Saturation of RNA Polymerase with DNA

Two sets of binding mixtures were prepared containing 18.8 pg or 8.4 pg of RNA polymerase per mixture, respectively. In each set, the enzyme was incubated with variable amounts of E. coli DNA (IO min, 37°C). The excess of free RNA polymerase was then inactivated by addition of heparin (50 fig in 10 ~1). Substrate solution was added to the mixtures 2 min later to start transcription. After a 10 min period of RNA synthesis, the reactions were stopped. The total RNA products and rRNA products were measured. “C-labeled rRNA (30.125 cpmlpg) was included in the hybridization mixtures as an internal standard. The values in the figure are averages of duplicate determinations, showing the amounts of 3H-UMP incorporated per reaction mixture into rRNA: (0-O) represents the incorporation in reaction mixtures with 16.8 fig of RNA polymerase; (O-O) represents the reaction mixtures with 8.4 pg of RNA polymerase. (x---x) shows the fraction of rRNA (% total RNA products) calculated for the reaction with 8.4 rg of RNA polymerase.

sheared DNA suggest that the viscosity of the solution may have a role.

Since this effect might possibly have lowered the apparent limit of template saturation, the weight ratio of enzyme/DNA was varied in another experiment by changing the concentration of RNA polymerase while keeping the DNA concentration constant. From the results of this experiment, shown in Figure 6, it is concluded that complete saturation of the template occurs at a weight ratio (enzyme/ DNA) of approximately 4 (arrow in Figure 6).

It should be noted that the fraction of rRNA transcripts is nearly constant at all DNA concentrations (see broken line in Figure 5). If rRNA promoters would differ from the other promoters merely in the rate constant of polymerase binding, it would be expected that the fraction of rRNA transcripts decreases with increasing ratio of RNA polymerase/ DNA. However, it remains constant near 15% even in the presence of a large excess of RNA polymerase over binding sites. This implies that preferential binding of RNA polymerase to rRNA promoters is the consequence of a high binding capacity of these promoters.


100

t30

60

_

LO

20

Amount of RNA pol (pg/reactlon mixture)

Figure 6. Saturation of E. coli DNA with RNA Polymerase

4.25 pg of E. coli DNA were incubated for IO min in binding buffer (3PC) with different amounts of RNA polymerase. Free enzyme molecules were then inactivated by treatment with heparin (50 rg per mixture; 2 min, 37°C). Transcription was allowed to proceed after addition of substrates (spec. Act. of 3H-UTP 1 .O Cilmmol) for 10 min at 37°C. Incorporation of 3H-UMP into rRNA (A-A) and nonribosomal RNA (O-O) were determined. The arrow indicates the assumed point of template saturation.

Number of RNA Chains Initiated by Heparin- Resistant Complexes Attempts to measure the number of RNA molecules initiated at rRNA promoters by analysis of chain- terminal nucleotides have not yielded unambigu- ous results so far. Thus the number of rRNA transcripts was estimated forming the ratio (total nucleotides incorporated into rRNA/number average chain length of rRNA). Sedimentation analyses of transcripts were used to determine number average chain lengths.

3H-adenosine-labeled transcripts were synthesized with heparin-resistant RNA polymerase-DNA complexes formed under conditions of template saturation. The DNA concentration was kept low enough to avoid the inhibitory effect of high DNA concentrations (see Figure 5). The transcripts were then centrifuged through sucrose density gradients prepared according to Kurland (1960). Figure 7 shows the distributions of ribosomal and total RNA products in twelve gradient fractions. The broken line essentially represents a number distribution of the rRNA transcripts. The broad range of molecular sizes is (qualitatively) expected for transcription at low ionic strength where the enzyme is known to terminate synthesis with a constant probability per unit time almost from the beginning of the reaction (Mueller and Bremer, 1969). The number average chain length (5) of total or ribosomal transcripts can now in principle be obtained from these distributions according to the relation

10 20 Sedlmentatlon coefflclent 1s)

Figure 7. Sedimentation of Transcripts

32 pg of RNA polymerase were incubated for 10 min at 37°C with 6.6 pg of E. coli DNA (20.2 nmol of nucleotides) in binding buffer (350 ~1). Excess RNA polymerase was then inactivated by addition of heparin (100 &g in 20 ~1). RNA synthesis was started 2 min later by adding 40 ~1 of a substrate solution containing JH-ATP (13.6 nmol. 29 &i/nmol) and the nonradioactive triphosphates (46 nmol each). The reaction was stopped after 10 min at 37°C by addition of 100 ~1 of 2.5% SDS, 0.1 M EDTA (saturated with diethylpyrocarbonate). 150 PI of the mixture were layered on top of a 13-ml sucrose gradient (20-5% sucrose in 0.01 M Na-acetate, 0.1% SDS, 0.1 M NaCI, 1 mM EDTA (pH 5.0)]. This gradient and a parallel gradient with ‘.C-rRNA as sedimentation marker were centrifuged for 16 hr at 27,000 rpm in an SW40 rotor of a Beckman ultracentrifuge (20°C).

Triplicate hybridization tests were performed with 200 (.LI samples of each fraction by incubating them with 250 ~1 formamide, 50 ~1 30 x SSC, and filters (100 rg of DNA) which had been preincubated with or without 30 rg of “blocking” rRNA. The amounts of competed radioactivity (A cpm) (O-O) and the ratios A cpm/chain length ( 0- - - 0) are plotted against the sedimentation coefficient. The chain length I, (nucleotides) was estimated for each fraction from the S value according to Kurland (1960). The radioactivity of the total transcripts in 200 ~1 of each gradient fraction (x-x) was measured by precipitation of small aliquots.

where n, = amount of nucleotides of a given class of RNA in fraction f, and If = chain length of the RNA molecules in fraction f, which may be determined from the sedimentation rate according to Kurland (1960). The values of n, may be calculated from the amounts of radioactivity in single gradient fractions (rr), from the specific radioactivity (a), the

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126

mole fraction of the labeled nucleotide (m), and the efficiency of hybridization (h):

rf nf a.rn.h

However, since the value of h was found in a control experiment to be independent of the position of the fraction in the gradient, and since the values of a and m can be assumed to be constants as well, i is calculated more conveniently according to a relation obtained by combination of equations (1) and (2):

i = g rf/i (rf/lf) From the data of Figure 7, the number average chain length of rRNA transcripts was found to be 970 nucleotides and that of total transcripts, 700 nucleotides.

The most critical source of error in this determination is a possible artificial fragmentation of transcripts. It may be noted here that no contaminant RNAase activities could be detected by addition of ‘YXabeled standard rRNA to parallel reaction mixtures and gradients. The validity of the method was tested by an independent determination of the number average chain length of total transcripts based on nucleotide analysis. Table 1 shows the nucleotide composition of 3H-adenosine- and 3H- guanosine-labeled transcripts which were synthesized in reaction mixtures parallel to those of Fig- ure 7. From these data, a number average chain length of 824 nucleotides is computed for total transcripts (see legend to Table 1). This estimate is only slightly higher than the value of 700 derived from the sedimentation pattern. It is expected that the sedimentation analysis should yield a some- what lower estimate than the nucleotide analysis, even in the absence of RNA breakdown. This is due to the fact that RNA molecules diffusing in the gradient to positions of lower S values are regis- tered by the application of the logarithmic relation between S value and chain length in higher numbers than the molecules diffusing to positions of higher S values. This systematic error should be largest in the uppermost fractions of the gradients. If the RNA fraction with the lowest S value in Figure 7 is disregarded for the calculation of the number average chain lengths, values of 1140 or 960 nucleotides are obtained for rRNA or total transcripts, respectively.

Aside from this systematic error, the question should be considered whether the results of the two methods of chain length determination are ac- tually comparable. It was shown previously by Mueller and Bremer (1969) that the chromato- graphic isolation of transcripts used for the nucleo-

tide analysis recovers also RNA chains which are too short to be acid-precipitable. We find also that some of the transcripts synthesized under the conditions of the experiment of Figure 7 are not precipitated by acid. These molecules do not contribute significantly to the total weight of the RNA, but they usually contain 20-40% of the 5’ termini. In con- trast to the nucleotide analysis, the sedimentation analysis of the total transcripts described above is not expected to record nonprecipitable RNA molecules which are presumably present in the top fractions of the gradient. It is conceivable, then, that the apparent agreement of the two chain length determinations results from a systematic error of the sedimentation analysis which compensates the failure of the method to record nonprecipitable transcripts. The magnitude of this possible error is not known. However, since the number of nonprecipitable RNA molecules may represent up to 40% of the total transcripts, the compensating error will probably not exceed 40%. This figure may serve to define upper limits of the number average chain lengths. They are 1140/0.6 = 1900 nucleotides for ribosomal RNA, and 960/0.6 = 1600 nucleotides for nonribosomal RNA.

On the basis of these values and of the total

Table 1. Nucleotide Analysis of RNA Produced by Heparin-Re- sistant RNA Polymerase-DNA Complexes

Amounts of Nucleotides (pmol per Reaction Mixture) in:

Nucleotide Total Transcripts

AP 1642

GP 2172

Total Xp 6242

PPPAP 5.2

PPPGP 4.8

i 624

rRNA

194

772

Parts of the reaction mixture of Figure 7 were used to measure the incorporation of 3H-adenosine into rRNA transcripts (duplicate hybridization tests) and into chain terminal and internal positions of total RNA (Experimental Procedures). Analyses of chain termini were also performed with 3H-guanosine-labeled transcripts which had been synthesized parallel to the ‘H-adenosine-labeled transcripts. The reaction mixture contained 3H-GTP (20 nmol) at a specific radioactivity of 15 &i/nmol.

The total nucleoside monophosphates (“total Xp”) incorporated into total transcripts were estimated from the amounts of AMP and GMP (“Ap,” ” Gp”) assuming the molar ratio of purines/ pyrimidines to be 0.95 (Chamberlin and Berg, 1962). For the calculation of a total “Xp” in rRNA, the mole fraction of AMP was taken to be 0.251 (Bolton. 1967). The number average chain length of total transcripts (i) was calculated by dividing the total amounts of monophosphates by the total amounts of nucleoside tetraphosphates derived from 5’ terminal positions (“pppAp,”

“PPPGP”).


nucleotide incorporation (“total Xp” in Table l), it can now be calculated that at least 8242/1600 = 5.15 pmol of total RNA chains were initiated on 6.6 pg of polymerase-saturated template, including 772/1900 = 0.41 pmol of ribosomal RNA molecules. Taking the molecular weight of the E. coli genome as 2.8 x log (Cairns, 1963), we estimate that at least 175 ribosomal and 2000 nonribosomal RNA chains were synthesized on a genome equivalent of DNA.

The Binding Capacity of rRNA Promoters To determine the average number of rRNA transcription units per genome equivalent, E. coli DNA was hybridized with saturating amounts of 3H- rRNA. From the maximum amount of 3H-rRNA bound to a given weight of DNA (details not shown), it was calculated that a fraction of 0.33% of the DNA used for binding reactions is complementary to rRNA. This result agrees well with other reports (Yankofsky and Spiegelman, 1962; Yu, Ver- meulen, and Atwood, 1970; Pace and Pace, 1971). A genome equivalent of DNA (2.8 x IO9 daltons or 8.3 x lo6 nucleotides) thus contains 27,400 nucleotides in sequences complementary to rRNA. Since in bacteria the rRNA transcription units contain one gene for each rRNA species and a single promoter region proximal to the 16s rRNA gene (Pato and Meyenburg, 1970; Bremer and Berry, 1971; Doolit- tle and Pace, 1971), the average number of transcription units per genome equivalent is calculated by dividing the total chain length of transcribed rDNA (27,400 nucleotides) by the summed chain lengths of 16s and 23s rRNA (4700 nucleotides, estimated from molecular weights of rRNA species given by Attardi and Amaldi, 1970). There are on the average 5.8 transcription units in a genome equivalent of the template.

It was concluded from the data of Figure 7 and Table 1 that the rRNA promoters in a genome equivalent of DNA protect at least 175 RNA polymerase molecules. This corresponds to a binding capacity of a single rRNA promoter region of 175/ 5.8 = 30 RNA polymerase molecules.

Discussion

The rRNA genes of E. coli represent only 0.6% of the genome, yet they support approximately half the total rate of RNA synthesis in vivo (Mueller and Bremer, 1968; Salser, Janin, and Levinthal, 1968; Lazzarini and Dahlberg, 1971). Compared with the average transcriptional activity of the rest of the genome, the rRNA genes thus exhibit a more than 100 fold higher activity under optimal conditions of cell growth. These differences are not accounted for by differences in the rates of RNA chain growth

(Dennis and Bremer, 1973) but reflect different promoter efficiencies in chain initiation. “Preferential transcription” of rRNA genes was also observed in vitro, in agreement with previous reports (Hasel- tine, 1972; Pettijohn, 1972; Birnbaum and Kaplan, 1973; Travers et al., 1973; Travers, 1973; Van Ooyen et al., 1975), although in vitro the transcription rate of rRNA genes was found to be only 15-30 fold higher than the average transcription rate of genome segments coding for nonribosomal RNA. This comparatively low preference of RNA polymerase for rRNA synthesis in vitro does not mean by itself that the system is lacking an element of posi- tive control of rRNA promoters; it may also be explained by a relatively high rate of initiation of nonribosomal RNA chains under in vitro conditions (Van Ooyen et al., 1975).

The experiments reported here provide evidence concerning the in vitro interaction of RNA polymerase with rRNA promoters. The measurements of specific RNA polymerase binding were based on the fact that the RNA polymerase holoenzyme forms tight complexes with promoters and thereby becomes insensitive against polyanionic inhibitors such as heparin (Zillig et al., 1970), poly(dAT) (Mueller, 1971), or polyinosinic acid (Bautz, and Beck, 1972). We chose heparin to discriminate between free and promoter-bound enzyme, mainly because of its technical convenience. The heparin- resistant RNA polymerase-DNA complexes are equally insensitive against the other polyanionic inhibitors (C. Oebbecke, unpublished data) and homogeneous with respect to their stability.

If preformed RNA polymerase-DNA complexes are allowed to transcribe the template in the presence of heparin, the fraction of rRNA molecules among the transcripts is found to be as high as in the absence of heparin. This indicates that the preferential transcription of rRNA genes is established during complex formation and does not primarily reflect a high intrinsic rate of rRNA chain initiation. Udvardy, Sumegi, and Venetianer (1974) proposed that the high efficiency of rRNA promoters might be related to their ability to bind RNA polymerase very tightly. This seems improbable in view of the fact that the activities of promoters of phage fd are not correlated with the dissociation rates of their complexes (P. H. Seeburg and H. Schaller, submitted for publication). The question may then be asked whether rRNA promoters differ from other E. coli promoters in the rate constant of polymerase binding or in the binding capacity (number of binding sites per promoter). Our results favor the second alternative, because the degree of preference of polymerase binding to rRNA genes was found to be almost constant at all weight ratios of RNA polymerase/DNA, even at large excess of enzyme mole-

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cules over binding sites. However, the two alterna- tives do not have to exclude each other completely (for instance, if rRNA promoters would comprise heterogeneous binding sites as suggested by Travers, 1975).

Ribosomal RNA precursor molecules are transcribed in bacteria from transcriptional units containing one gene each for 16s rRNA, 23s rRNA, and 5s rRNA with a promoter region proximal to the 16s rRNA gene (Pato and Meyenburg, 1970; Bre- mer and Berry, 1971; Doolittle and Pace, 1971). It was calculated that at least 30 RNA polymerase molecules per rRNA transcription unit can be protected from heparin by complex formation with E. coli DNA. This result may be taken to indicate that a rRNA promoter has the capacity to bind 30 RNA polymerase molecules if heparin resistance is assumed to be a valid criterion of promoter-bound enzyme (Schafer et al., 1973b). The number 30 is similar to the number reported for the binding capacity of “promoters” of phage T5 by Schafer et al. (1973a). However, it is higher by a factor of 7 than the binding capacity of rRNA promoters of E. coli determined by Venetianer et al. (1976) who fol- lowed the same experimental approach. The number of 4-5 RNA polymerase molecules per promoter reported by these investigators certainly represents only a lower limit, since it was calculated from the weight ratio of rRNA transcripts/rDNA on the assumption that the transcripts had the full length of the rRNA precursor. Taking into account the fractions of shorter RNA molecules shown in the sedimentation profiles of Venetianer et al. (1976), we estimate that according to their data, the binding capacity of rRNA promoters should be 6-10 RNA polymerase molecules. Our results still differ con- siderably from this figure. The discrepancy is found in the absolute numbers of RNA polymerase molecules protected from heparin by a given amount of DNA, not in the relative amount of rRNA promoter complexes. It can be excluded that the higher binding capacity measured in our system is due to insufficient heparin treatment (see Figures 1 and 2). The value of 30 RNA polymerase molecules per promoter is a conservative estimate, taking into account different sources of errors of the sedimentation analysis. It therefore seems most probable that the quantitative discrepancy between our results and those of Venetianer et al. (1976) arises from differences in the reaction conditions rather than in the techniques of measurement. For instance, it is possible that the 4 fold higher Mg++ concentration used by Venetianer et al. (1976) re- stricts the binding capacity of the promoters as shown in the case of the lac and gal promoters by Nakanishi et al. (1975).

Even if the evidence for multiple RNA polymerase

binding to rRNA promoters is accepted from a technical point of view, one should be aware of the possibility that this phenomenon could be an arti- fact of the in vitro conditions, or that the criterion of heparin resistance is not strictly specific for promoter-bound enzyme. It may be argued that some of the enzyme molecules protected from heparin are initially bound to nonspecific template sites and drift during the transcription periods of binding tests into promoter regions. But if this were the case, there would not be any obvious explanation of the fact that the numbers of heparin-resistant polymerase molecules per promoter show well defined saturation values which are widely different for different promoters (Schafer et al., 1973b; Will- mund and Kneser, 1973). It would be desirable to test the postulate of multiple RNA polymerase binding using criteria other than heparin resistance for measurements of RNA polymerase-promoter complexes.

A possible significance of multiple polymerase binding cannot be dismissed at this point. It is plausible that variations of promoter efficiencies could have evolved by variations of the number of RNA polymerase binding sites within promoter regions. Multiple binding sites may increase the ability of promoters to compete with other promoters for free, active RNA polymerase molecules which, according to Bremer and Dalbow (1975), exist in bacterial cells at limiting concentration. They would also allow several primary polymerase-promoter complexes simultaneously to be converted into “open complexes” and thus be prepared for rapid chain initiation.

Most or all of the binding sites of rRNA promoters could conceivably function also as initiation sites. However, the slow initiation of rRNA transcripts suggested by the data of Figure 3 would be explained most easily by the assumption that the RNA polymerase molecules lined up in a binding region must drift to a single initiation site to start transcription.

If the efficiency of promoters is a function of the multiplicity of RNA polymerase binding, one would predict that most of the 2000 enzyme molecules accomodated by promoters other than rRNA promoters are bound singly or in small clusters. The fact that nonribosomal RNA synthesis proceeds at the highest rate already at the beginning of the reaction agrees with this proposition.

Experimental Procedures

Chemicals Heparin (pharm., USP XVII), sucrose (RNAase-free), and nonradioactive ribonucleoside triphosphates were obtained from Serva (Heidelberg). Rifampicin was from Boehringer (Mannheim). DNAase (deoxyribonuclease 1) and RNAase (ribonuclease A) were products of Worthington (New Jersey). The RNAase stock solution


was heated before use at 90°C for 10 min. 2-3H-adenosine-5’- triphosphate (27 Ci/mmol). 6-3H-guanosine-5’-triphosphate (15 Ci/mmol), 5-3H-uridine-5’-triphosphate (20 Ci/mmol), 6-3H-uracil (24 Ci/mmol), and 2-“C-uracil (59 mCi/mmol) were purchased from Amersham Buchler (Braunschweig). Formamide (p.A.) was from Merck (Darmstadt).

Preparation of E. coli DNA E. coli B cells were grown in glucose minimal medium (generation time 65 min). DNA was isolated from the cells according to Miura (1967) and dialyzed against 0.05 M Tris-HCI (pH 7.2). 0.1 M NaCI, 1 mM EDTA, or binding buffer (see below). Sedimentation analysis (Burgi and Hershey, 1963) indicated an average molecular weight of 6.3 x IO’. The concentration of DNA preparations was determined by spectral analysis as described by Felsenfeld (1968). Weights were calculated on the assumption that E. coli DNA nucleotides have an average molecular weight of 335 daltons (monosodium salt).

Preparation of RNA Polymerase DNA-dependent RNA polymerase was isolated from frozen E. coli B cells (Merck, Darmstadt) as described previously (Mueller, 1971). except that an additional step of sedimentation through a glycerol gradient [30-10% glycerol in 10 mM Tris-HCI (pH 7.9), 25 mM KCI. 5 mM MgCIZ, 0.1 mM dithiothreitol] was performed. The enzyme was stored in glycerol gradient medium at -70°C.

Preparation of Ribosomal RNA E. coli B cells were grown in glucose minimal medium with or without addition of radioactive uracil. At a cell titer of 5 x 108. rifampicin (50 wg/ml) was added, and aeration at 37°C was continued for IO min. The cells were harvested by centrifugation, suspended in 10 mM Tris-HCI (pH 7.8). 10 rr!M Mg-acetate, and lysed by a French pressure cell. The homogenate was treated with DNAase (20 Kg/ml, 5 min. 37°C). cleared of debris by centrifugation, and adjusted to 0.5 M NH&I. 10 ml portions were layered onto IO ml cushions of 15% sucrose in 10 mM Tris-HCI (pH 7.6), 10 mM Mg-acetate, 50 mM NH&I, and centrifuged in a Beckman 50.1 rotor at 40,000 rpm at 4°C for 2.5 hr. The ribosomal pellets were suspended in “low Mg buffer” [lo mM Tris-HCI (pH 7.9), 0.2 mM Mg-acetate]. The suspensions were then centrifuged on 3 ml cushions of 15% sucrose in the same buffer (rotor SW5OL, 40,000 rpm, 4”C, 6 hr). The pelleted ribosomal subunits were taken up in “low Mg buffer” and dissociated by incubation with 1 vol of 1% sodium dodecylsulfate (SDS), IO mM EDTA at 37°C for 5 min. The rRNA species were separated by sedimentation through a sucrose gradient [30-5% sucrose in 10 mM Tris-HCI (pH 7.6). 1 mM MgCI,, 0.1 M NaCI, 0.5% SDS; 36 ml) which was centrifuged in an SW 27 rotor at 27,000 rpm at 20°C for 14 hr. Fractions containing 16s and 23s rRNA were pooled and shaken with 1 vol of phenol [saturated with SSC; SSC = 0.15 M NaCI, 0.015 M Na-citrate (pH 7.0); n x SSC = n fold concentration of SSC]. The RNA was precipitated from the aqueous phase after addition of 2 vol of cold ethanol, collected by centrifugation, and dissolved in 2 x SSC. The concentration of rRNA preparations was determined from the absorb- ance at 260 nm and neutral pH (A,., x 10m3 = 8.07).

Conditions of RNA Polymerase Binding and RNA Synthesis Reaction mixtures were usually prepared by adding 16.8 pg of RNA polymerase (in 20 &I) to 200 ~1 of a solution containing 6.5 pg of E. coli DNA in binding buffer [lo mM Tris-HCI (pH 7.9), 50 mM KCI. 5 mM MgCI,, 0.05 mM EDTA, and 0.5 mM dithiothreitol]. Afler incubation for 10 min at 37°C. 50 pg of heparin were added (in 10 ~1). and the incubation was continued for 2 min. Heparin- resistant RNA polymerase-DNA complexes were then allowed to start transcription by adding 300 ~1 of a substrate solution which contained 0.02 mM SH-UTP (0.5 Ci/mMol), 0.4 mM ATP, 0.4 mM GTP, 0.4 mM CTP. 60 mM Tris-HCI (pH 7.9), 10 mfvl MgCI,, 0.1 mM EDTA, and 0.1 mM dithiothreitol. After a 10 min period of RNA synthesis at 37”C, the mixtures were shaken with 600 ~1 of phenol

(saturated with 2 x SSC) and 100 ~1 of 14 x SSC. The aqueous phase (“transcript solution”) was used for analysis of the RNA products. Deviations from this standard procedure are indicated in the legends to the figures.

Measurement of rRNA Transcripts by Hybridization E. coli DNA was incubated in 0.33 M NaOH at 37°C for 90 min, and after neutralization and adjustment to 6 x SSC, adsorbed to nitrocellulose filters (Sartorius, Gbttingen) by filtration. The filters, each containing 100 fig of DNA, were left to dry at room temperature. They were then heated at 60°C for 2 hr and used for RNA-DNA hybridization under conditions which were essentially those of Gillespie and Gillespie (1971). The filters were placed singly into vials of two sets: vials of one set containing nonradioactive rRNA (20-30 rg) in 0.4 ml formamide solution (50% formamide, 3 X SSC). and those of the other set containing 0.4 ml of formamide solution without rRNA. After 24 hr of incubation at 37°C. the filters of the two sets (having “blocked” or “nonblocked” rDNA sequences, respectively) were rinsed with 50% formamide. 3 x SSC. and transferred into vials containing 250 ~1 of formamide. 50 ~1 of 30 x SSC. and 200 ~1 of transcript solution. Hybridization of radioactive RNA products was allowed to proceed for 46 hr at 37°C. The filters were then washed twice with cold 2 x SSC (5 ml per filter), treated with RNAase (20 pglml in 2 x SSC) for 50 min at room temperature, washed again twice by suspension in 2 x SSC. dried, and placed into vials with scintillation fluid (toluene with PPO and dimethyl-POPOP at concentrations of 3.64 g/l or 0.36 g/l, respectively).

The radioactivity in the rRNA fraction of transcripts was determined from the difference of the radioactivities found on filters with “blocked” and “nonblocked” rDNA (A cpm) and from the hybridization efficiencies of standard rRNA (isolated from ribo- somes) which was used as an internal or parallel reference: cpm in rRNA = A cpm/H. - H,,. H, and H. designate the fractions of standard rRNA hybridized to blocked (HJ or nonblocked (H.) rDNA. The value of H. was on the average 0.65 and was nearly independent of the rRNA input within the range of l-50 ng rRNA per filter. The value of Hb was usually < 0.03. The average yield of hybrids of nonribosomal RNA products at inputs of approximately 10 ng of RNA per filter was 35%. Part of the nonradioactive “blocking” rRNA bound to the filters after the first hybridization step was found to be only loosely associated with the DNA. Re- moval of this fraction (corresponding to approximately 1% of the DNA) by RNAase treatment or extensive washing in formamide solution at 37°C did not significantly affect the measurement of rRNA transcriots.

Measurement of Total RNA Transcripts Aliquots of transcript solutions were mixed with 2 ml of a solution of cold carrier RNA (yeast RNA, 50 Kg/ml, in 2 M NaCI) and 0.5 ml of 3 M trichloroacetic acid. The mixtures were kept in ice for 2-10 min; they were then filtered through nitrocellulose filters (Sarto- rius, GOttingen; 0.45 pm pore size). The filters were washed 4 times with dilute trichloroacetic acid (approximately 5 mM), dried, and counted in scintillation fluid. The counting efficiency of tritium radioactivity in precipitated RNA was equal to that of hybridized RNA (27%).

Analysis of RNA Chain Terminl Hybridized RNA or RNA products isolated from reaction mixtures by adsorption to DEAE paper according to Mueller and Bremer (1969) were hydrolyzed in 0.4 M KOH (16 hr, 37°C) and analyzed as described in detail previously (Mueller and Bremer, 1969). The nucleosides and nucleotides derived from chain terminal and internal positions were separated on paper strips (Whatman 3MM) by high voltage electrophoresis. To measure the distribution of radioactivity in the paper strips, 1 cm sections were placed into vials with 6 ml of a solution composed of 6.5 vol of scintillation fluid (see above), 1.5 vol of Biosolve BBS3 (Beckman), and 1 vol of 0.2 M NaCI. The vials were shaken at room temperature for at least

Cell 130

2 hr. and the radioactivity was counted without removing the paper sections from the vials. Under these conditions, the counting efficiency of tritium radioactivity in nucleotides was 24.6%.

Acknowledgements

This paper is dedicated to Professor 6. Rensch on the occasion of his 75th birthday. This work was supported by the Deutsche Forschungsgemeinschaft.

Received September 13, 1976; revised October 18. 1976

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Capacity of ribosomal RNA promoters of E. coli to bind RNA polymerase

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