S4-α mRNA translation regulation complex

.J. Mol.

Hiol.

(1987)

196. 323X332

Sk

mRNA

Translation

Regulation

Complex

II. Secondary Structures of the RNA Regulatory Site in the Presence and Absence of S4 Ingrid C. Deckman? and David E. Draper1 Department

of Chemi.stry

Johns Hopkins University Baltimore, MD 21218, 1Y.S.A. (Received 30 September 198,

and in revised form 23 February

1987)

The secondarv structure of the Escherichia coli tl mRNA leader sequence has been determined uiing nucleases specific for single- or double-stranded RNA. Three different length c1RNA fragments were studied at 0°C and 37°C. A very stable eight base-pair helix forms upstream from the ribosome initiation site, defining a 29 base loop. There is evidence for base-pairing between nucleotides within this loop and for a “pseudo-knot” interaction of some loop bases with nucleotides just 3’ to the initiation codon, forming a region of complex structure. A weak helix also pairs sequences near t)he 5’ terminus of the c1mRNA with bases near the Shine-Dalgarno sequence. Affinity constants for the translational repressor S4 binding different length c( mRNA fragments indicate that most of the S4 recognition features must be contained within the main helix and hairpin regions. Hinding of S4 to the c1mRSA alters the structure of the 29 base hairpin region, and probably melts the weak pairing between the 5’ and 3’ termini of the leader. The pseudo-knot structure and the conformational changes associated with it provide a link between the structures of the S4 binding site and the ribosome binding site. The u mRNA may therefore play an active role in mediating translat’ional repression,

1. Introduction Regulation

of ribosomal protein synthesis in coli takes place at the translational level by an autogenous mechanism. In the case of the x operon. the ribosomal protein 54 is both encoded by the a operon and bound at a target site within the c( mRNA leader sequence to repress translation. S4 also binds to the 16 S rRNA to initiate ribosome assembly. In the accompanying paper (Deckman et al., 1987), we showed that most of the 84 recognition features are contained within bases 23 to 69 of the TVpromoter transcript. Known ribosome recognition features span bases 74 to 107 (Gold et al., 1981). but’ S4 binds too weak]? in this region to compete well with ribosomes. With t’he idea that the CImRh’A secondary st’ructure might provide a connection between the 54 and ribosome binding sites. we have used structure mapping techniques to investigate the mRSA structure in the presence Escherichia

and absence of S4. The results show t,hat the main S4 binding region is linked to bases just 3’ to the initiation codon. A distinct change in mRNA conformat)ion is induced by S4 binding, and could be important in mediating translational repression.

2. Materials and methods (a) Reagents md buffm

All buffers were prepared tising water run through a Millipore MilliQ deionizer and filter. Stock solutions were autoclaved to prevt‘nt nurlease contamination. Salts and buffers were standard reagent grade. TMK buffer contains 0.35 M-KU. 20 mlcl-Tris. HCI (pH 7.6). 20 mMMgCI,. TK buffer is the same, but with no MgCI,. [y-3~I’]A’IT (3000 Ci/mmol) was purchased from Amersham. T,. T,. and V, nuclrases were purchased from Roehringer-Jlannheim. I3RL, a11d Pharmacia-P-I,. rrsprctivrly.

(b) Prepration t Present address: Merck, Sharp, & Dohme Laboratories. $ Author addressed.

Research West Point. PA 19486, U.S.A. to whom correspondence should hc

oj 32P-lahrlcd

KiV.4 s

Cnlabeled RXAs were prepared hj in-vitro transcription as described in the accompanying paper. After 1 h incubation at 37°C’. a 250@ transcription reaction was run immediately over a 5.5 cm Superose 12

c*olumn (Pharmacia) in 20 m.n-Tris. HCI (pH 7.6). (1.1 ;\I?ja(‘l buffer at 0.6 ml/min to separat)e full-length REA from unincorporated nucleoside triphosphates. tc~mplat(~ IlKA fract,ion was and smaller RBAs. The peak precipitated with ethanol and 20 pmol t,reated with bac%erial alkaline phosphatase (BRL) for 20 min at 45’C in 10 mm-Tris. HCI (pH 7.6). 1 mM-EDTA. Prot,einase K. (Boehringer-Mannhrim) was added to a final c.oncZentration of 0.1 mg/ml and the reaction incubated for I h at 37°C to destroy any phosphatase acltivitv. After extractions with phenol and ether and precipitation with rthanol. most R?u’As were ready to be labeled, We found that the largest R?I;A labeled more efficiently if first purified by elertrophoresis in a 50”,, (w/v) urea/6(+, (w-/v) acrylamide gel. 5’ end-labeling with phage T4 polynucleotide kinase (I’.S. Riochemicals) was carried out with 20 pmol of RKA fragment in IO ~1 with 50 mM-Tris. HCI (pH 7.6), 10 mMMgSU4, 5 mCi [y-3ZP]ATP/ml. After incubation at 37°C for 40 min, the reaction was loaded directly on to a fiO?b urea/6Oi, acrylamide gel, t’he wells of which had been soaked for 10 min with 1.4 M-/-mercaptoethanol to minimize radiation damage. The band of RNA was detected by autoradiography, excised with a sterile razor blade. and extracted by the “freeze-and-squeeze” met,hod described by Kean & Draper (1985). The RNA was precipitated using yeast tRNAPhe (Sigma) as carrier. (c) RNA structure mapping experiments Dilutions of ribonucleases T, , T, and V, were prepared in a buffer containing 25% (v/v) glycerol, 5 mM-Tris HCI For the S4 protection (pH 7+), 05 mM-EDTA. experiments, 1 ~1 of purified 84 (in buffer with 6 M-urea) or 1 ~1 of 6 M-urea (for a control) was incubated in 20 ~1 of TMK buffer for 30 min at 37°C. To 5 ~1 of the renatured protein (or 5 ~1 control) was added I ~1 of labeled RNA (20,000 cts/min) in TK buffer. Incubation was continued at 37°C for 10 min to allow R’NA renaturation and protein binding. (The RXA could not be renatured in TMK buffer at 65”C, as standard for filter binding assays, because a small percentage of the molecules hydrolyze and give an unacceptable back-

ground on sequencing gels. Control experiments showed that this renaturation protocol gave the same binding as found previously.)

Reactions

were cooled to the desired

temperature, and 1 ~1 of the indicated nuclease dilution was added and allowed to react for 5 min. Reactions were quenched with 5~1 of buffer containing 10 M-urea, 0.00257& (w/v) bromophenol blue and xylene cyanol, and 2.5 mM-EDTA, and run on an 85 cm 504, urea/g% acrylamide gel at 40 W, as described (Kean & Draper, 1985). The smallest RNA was sometimes run on 40 cm, 12% acrylamide gels. Autoradiography was for 24 h with pre-flashed film and an intensifying screen.

3. Results (a) Strategy

of the structure

determination

To investigate the structure of the c1mRNA leader we have carried out “structure mapping” experiments (Wurst et al., 1978); i.e. we determined the susceptibility of each base in the sequence to nucleases specific for helical or single-stranded RNA. The RNAs used in this study were all transcribed from plasmids containing phage T7 RNA polymerase promoters, and had 25 bases at

the 5’ t’erminus t,hat arrl not contained in trarrscaripts from ihe GIpromot,er (Deckman it al., 1987). Thrc>ck different RNA fragments were ustd. Thts I?. coli sequence in each begins in the xpc operon I6 nucleotides upstream from the u promoter start site (Cerretti et al., 1983): while the 3’ tttrminus is located eit’her at C204 of t*he CIpromoter transcript, at’ Cl01 or at U69. The shortest fragment lacks all the feat,ures known to be required for ribosomr initiation. S4 binds to the longest fragment with the greatest affinity (13 pb1-l): the affinity decreases 1)~ about a factor of 2 with each successive d&&on (Deckman et al., 1987). For convenience these three molecules will be referred to as (169. (‘101 or (‘204 RNA. We used two enzymes specific for single-stranded RNA, T, and T, RNase. T, RNase cleaves only on the 3’ side of G residues, while T, is not base specific, but does prefer to cleave 3’ to A residues (Uchida bi Egami, 1967). Both enzymes leave 3’ phosphates after cutting. VI nuclease from cobra venom was also used (Lockard $ Kumar, 1981). This enzyme has a strong, but not absolute. specificity for nucleotides in Watson-(Irick helixes (Lowman & Draper, 1986). V, nucleasr leaves 5’.terminal phosphates. The affinity cleavage reagent

methidiumpropyl-EDTA-Fe(I1)

(Hertzberg

& Dervan, 1982) was also reacted with the CImRNA. This intercalator shows occasional tight binding to unusual RNA structures (Kean et al.. 1985), but did not give any useful information wit-h the o! mRNA. A potential problem with structure mapping experiments is that cleavage at one site may relax the RT\‘A structure in such a way that cleavage at a second. normally unreactive site, rapidly follows. This behavior has been detected in the 5 S rRNA 1)~ comparing the digestion patterns of RNAs labeled at the 5’ or 3’ ends (Garrett & (Heson, 1982). In most cases we were unable to use 3’ end-labeled RNA, since t’he T7 transcripts tend to be a mixture of two RNAs with 3’ termini differing b,v one ba,se (Draper et al., 1987). The exception was the (1101 RNA, and digestions of this RNA did appear to be identical with 3’ and 5’.labeled material. We attemptSed to distinguish primary from secondar? cutting sites by carrying out each reaction with three or four enzyme concentrations: and examining patterns for changes in relativtb the digestion intensit’ies as t,he extent of digestion when a high proportion of the

increases.

Onl>

molecules wer’ca cleaved ( > 30%) were changes in the digest’ion patterns detected. An example sequencing grl autoradiograph is shown in Figure 1. To use the struc%urr mapping data in a search for the correct z mRIvA systematic secondary structure, we had a computer search for all possible helices four base-pairs and longer. G . (‘, A. LT and G. I: pairs were assigned weights of 3, 2 and 1, respectively, and helices not reaching a minimum weight, (in 4 base-pairs) were filt,ered out of the display. The matrix of base-pairing possibilities in the a leader sequence is shown in

325

tl mRNA Structure

Figure 1. Autoradiograph of partial V, digestions of the Cl01 RNA run on an 85 cm denaturing acrylamide gel. Digestions were carried out at 0, 37 or 37°C in the presence of S4, as indicated; 3 digestions using different dilutions of nuclease were made in each case. The other lanes are labeled OH-. alkaline hydrolysis; T,, T, RNase digestion under denaturing conditions; C, control (no VI nuclease added).

Figure 2. A large number of weak potential pairings are found (filter set to a low value), while potential strong helices are much less frequent. We then superimposed our results on this matrix to find helices allowed by the data. G * A juxtapositions in helices, which are not uncommon in ribosomal RNA structures (Noller, 1984) and form hydrogenbonded pairs in DNA helices (Brown et al., 1986), were allowed as base-pairs in some of the computer searches. Only minor additions to the matrix were found. The a leader sequence was also searched with a computer program that uses the Nussinov & Jacobson (1980) algorithm and estimated base stacking free energies (Tinoco et al., 1973) to

I

20

I

I

40

I

I

60

I

I

80

I

I

construct, a matrix of base-pairing possibilities. The program is able to search the matrix for structures within a specified free energy interval (D. E. Draper, unpublished results). About 100 different structures generated by this program were considered for their fit to the data. In presenting our data, we have divided the molecule into three sections, which for the most part can be discussed individually. For each section we show the most likely secondary structures. A structure of the complete molecule is shown in Figure 9. to help place the sections in context. (b) Structure of the main helix

We find that there is only one helix that is 100 present unambiguously in all three c1mRNA 1 fragments under all conditions; we refer to it as the “main helix.” The helix pairs C23-11’31 with A61G68, and is shown along with nuclease cutting data in Figure 3. There are three main arguments for its existence: strong V, nuclease cutting extends along the 5’ strand of the helix; the only G residues in the 01leader completely resistant to T, RNase occur in this helix; and it is predicted to be a stable helix reasonable present in all thermodynamically secondary structures by both computer programs we have used (see Fig. 2). The V, nuclease digestion pattern shown in Figure 3 is identical in all three RNA fragments examined, with the minor exception of more intense cutting at US5 in the U69 RNA. A single base from the Us sequence at bases 25 to 27 must be bulged to form the helix. There is

23

U. - c G7J*U’C*C”AoUoI

Figure 2. Matrix of base-pairing possibilities in the leader sequence (Cl01 RNA fragment). For a possible base-pairing to be displayed, it had to be part of a 4 base-pair helix with a minimum weight of 10 (underlined) or 8. The weights assigned were G. C pairs 3, A . U pairs 2, and G. U pairs 1. G . U pairs are indicated by open circles. Numbers next to helices indicate the figure in which the helix is drawn.

66 -#,A

I

I

1

I

I

I

3I

I

A G G U,A

-61

a mRNA

Figure 3. Nuclease digestion data for the main helix region of t( mRNA. Numbering of bases is from the 5’ nucleotide of the transcript. Dots indicate sites of V, nuclease digestion; the size of the dot is approximately proportional to the rate of nuclease digestion. No T, or T, RKase cutting was observed in this region.

-.---.-_

fragment. new T, RNase cut,s appear at positions Cl6 to lr21 (Fig. 4(c)), while V, nucleasr cauts in thcl main helix are virtually unchanged. The new Tz RIXase cuts are consistent with U69 pairing with G22. The only other likely secondary structure that incorporates the set of strong V, cuts at (:24 to 1’33 in a helix pairs Cl9 to A35 with 1T7.5 to A94. This structure cannot form in the U69 RXA and would give a much different T, RNase digestion pattern in the longer R#NA fragments.

(c) Structure I

I I I I Ill I I 111I I IdiuG’cGI

~~~‘A’AUCUUUUGUAUGUC~U

I 16

(cl

.I

I

UGC69

Figure 4. Structure in the weak helix region of the CImRNA. Symbols indicating nuclease digestion sites are: dots, cobra venom V, nuclease; lines, T, RNase; wedges, T, RNase. The size of a symbol is approximately proportional to the rate of nuclease digestion. (a) and (b) Digestion data at 37°C (Cl01 RNA) superimposed on 2 different possible base-pairings. (c) Digestion data at 37°C for the U69 RNA. no known reason why all three possible single hase bulges not be should thermodynamically equivalent. We have drawn U26 as the bulge, since nuclease clearly skips the phosphodiester VI between U26 and U27 in cutting the helix. This interpretation is supported by studies of an rRNA helix containing a single base bulge at one of two consecutive A residues. Diethylpyrocarbonate reactivities indicate that the 5’ A is preferentially bulged (van Stolk & Noller, 1984). A series of V, nuclease cuts along this helix is interrupted only on the 3’ side of the bulged A (Kean & Draper, 1985). An observation that confirms the existence of the main helix is a comparison of T, RNase cutting in the U69 and C204 RNAs. Tn the shorter RNA

of the “weak”

helix

Once the base-pairing of the main helix is established, it is logical to look for pairing between the 5’ and 3’ termini of the C( leader. There are several possibilities, all of them with a high proportion of A. U and G. U base-pairs, and therefore only marginally stable. Two sets of V, n&ease cuts are seen at positions U6 to Cl9 and A66 to U73 (Figs 4(a) and (b) and 5), which is good evidence for helix formation. The V, cuts at 1;6 to U19 are not found in the U69 RNA (Fig. 4(c)), but are found in the two longer RNAs; therefore the complementary sequence for U6 to Cl9 must lie within A70 to ClOl. (Other V, cuts in the U69 RlVA are identical with those in the Cl01 RNA, indicating that the RNA has not rearranged to a new structure.) Two base-pairing schemes that are consistent with the V, digestion data are shown in Figure 4(a) and (b). Ext’ensive T, and Tz RNase digestion also occurs throughout this region and must be taken into account. Particularly striking is the set of bases in the 5’ sequence that is susceptible to both VI and T, nucleases (LJ6 to U19). The most likely explanation for this is that the helix is near its melting point,. and can be trapped by a nuclease in either a singlestranded or double-helical form. To test this, the nuclease digestions were repeated a,t 0°C to stabilize base-pairing. Data are shown in Figure 5, The V, cuts remain. with minor variation, while t,he T, cuts

VI G OHT2 T2

Figure 5. Gel autoradiographs

of V, nuclease and T, RNase digestions of the C204 RNA 5’ terminus at 0 “C and 37 “C G indicates a T, RNase digestion under denaturing conditions. Electrophoresis of 5’ end-labeled RKA was from right to left; the sequence should be read with the 5’ terminus at the left side of the Figure.

327

o[ mRNA Structure

U “A; .U A U c .G -C” .C - G AA u

;rfj c

P - u; Jl - A .G l U

U-% A-U

:I$

LY

G-C C A U A’&

G C C%% I 100

,I,1 9-G U A U A G G - G*A U C’A’U’A’A’G I 130

C :: ’ ?I C G G I 170

Figure 6. Structure mapping data at 37°C for sequences 3’ to the CIleader, in the C204 RNA. T, and V, nuclease digestion data is represented as for Fig. 4.

completely consistent with these structures. The V, nuclease cuts at A119 to U124 become stronger at O”C, relat,ive to other cutting sites in the molecule, probably because the predicted stability of the helix

is only -2.2 kcal at 25°C. The second, longer helix should be much more stable, AG z - 12.8 kcal. The sequence G97 to G102 shows no sensitivity to T, or T, nucleases, even though flanking sequences are sensitive,

and is cut

at two

positions

by V,

nuclease. These t.wo cuts also become more intense at 0°C. There are no sequences 3’ to the main helix that could account, for a weak structure forming at, this position. A47 to U53, which is within the hairpin loop defined by the main helix, can potentially form a stable helix with G97 to c‘103. This

possibility

is discussed

further

in the

next,

become nearly undetectable. In addition, T, RNase cutting at A77-A79 is much slower (relative to other cutting at other sites) at 0°C (not shown). The structure shown in Figure 4(a) probably accounts for the digestion data better, though the Figure 4(b) structure cannot be ruled out. The free energy of

section,

adding structure

-8.6 kcal relative to the unpaired 29-base loop. respectively).

(b) onto the end of the main helix

is only - I .6 kcal at 25”C, and for structure (a) it is -1.2 kcal (Tinoco et al.? 1973), so it is not surprising that the structure is partially melted at 37°C (1 kcal = 4.184 kJ). (d) Structure in the 5’13 coding region

Our structure mapping data with the C204 RNA extend x70 nucleotides into the coding region of the first 01operon gene, S13. Potentially, this sequence can form two stable stem-loop structures, shown in Figure 6. The nuclease digestion data are

(e) Structure of the hairpin

region

The main helix defines a hairpin loop from bases U32 to C60. Two base-pairing schemes, shown in Figure 7(a) and (b), are thermodynamically (Tinoco rules stabilities of - 8.2 and

likely

A third structure incorporates the possibility of a interaction with nucleoso-called “pseudo-knot” tides just 3’ to the initiation codon (Fig. 7(c))?. t A “pseudo-knot” is the pairing of bases in a hairpin or bulge loop with bases outside the loop in the same molecule; if a full turn of helix were formed a true topological knot would be created. Evidence fbr pseudoknot structures in 16 S rRNA (Noller, 1984), some mitochondrial and fungal introns (Davies et al., 1982) and viral ItNAs (PIeli et al., 1985) has been presented.

(a)

(b)

AGU G u

A u

AGU G

A

Figure 7. Possible secondary structure in the hairpin loop region of the a mRNA. (a) and (b) Different possible basepairings with structure mapping data indicated for the Cl01 RNA at 37°C as for Fig. 4. (c) Possible pseudo-knot structures.

I. (Y. Deckman

328

Three-dimensional models of both pseudo-knot structures shown can be constructed, but it is not’ known how to estimate the AG value for forming this kind of structure. Just the five C. G base-pairs would contribute - 18.2 kcal in a normal helix, hut this must be at least partially offset by distortions of helices in the structure and the entropy of closing an additional loop. The pseudo-knot structure is proposed to account for the observation that sequences near the initiation codon affect the structure of the hairpin region. At 0°C the VI nuclease cutting is quite different for the three different size RNA fragments, both in the extent of cutting and in the relative intensities of the cuts (Table 1 and Fig. 8). At 37°C the V, nuclease digest are very similar, though not identical. Tn particular, new V, cutting sites at G49 and C48 appear as the RNA is lengthened from U69 to Cl01 and C204 (see Table 1). In looking for sequences near Cl01 that might be interacting with the hairpin region, we noted that V, and T, n&ease digestion data suggest that G97 to U103 is involved in a structure, though there are no complementary sequences available for forming a simple hairpin (Fig. 6). Potentially, the Cl01 RNA can form five of the seven pseudo-knot base-pairs possible in C204 RNA. Removal of the pseudo-knot pairing in two steps would explain why C204, Cl01 and U69 RNA all appear to have different hairpin region structures at O”C, and why G49 and C48 become progressively sensitive to V, nuclease as the RNA becomes longer. A pseudo-knot structure also provides an explanation for another peculiarity of the structure mapping data. Nucleotides between G82 and C99 are completely unreactive towards V, nuclease and only very weakly reactive with T, or T, RNases. This RNA sequence must extend between the ends of a long helix to create the pseudo-knot; with the

and D. B. Draper

Table 1 17, digestion of the a mRNA hairpin 35 I

U42

oc

ooc

4;i I

-50 I

55 I

~IUIJUAOUAU(:CUGAAAA(:(:G(:(‘I~lJCU(’A A. 0 “C U69 Cl01 (~204 l3. 37 “C U69 Cl01 C204 c. 37”CfS4 Cl01 C204 D. O”C+SI C204

2110010035300000000000000000 4220020123252110010000220200 32200001 34321 100023001

210000

3220010044420000000000110100 4220020033330000010000110100 4220000022220000011001

110000

3220010012232110010000220200 4220000122231 11001 10001

10000

4220000122231110011000110000

Approximate relative intensities of Vi nucleasecutting at nucleotides U31 to A58 within the hairpin region. Since nucleotides U27 to U33 are cut about the same under all conditions, cutting intensities are reported relative to these sites. Intensities were determined by densitometer scanning of the autoradiographs.

weak helix formed, 18 nucleotides are available to extend over 26 base-pairs. This is possible, but might require the single-stranded backbone to sit fairly close to the rest of the RNA, where it would be sterically inaccessible to nucleases. The structure mapping data in the hairpin regin are somewhat ambiguous, because several nucleotides are sensitive to both V, and T, nucleases. For instance, the V, nuclease cuts seen in the main helix extend to base U33, while T, RNase cutting is detectable at U32. This kind of overlap is usually resolved when digestions are carried out at 0°C to stabilize the structure (this was the case with the weak helix region). Although more extensive V, nuclease cutting is seen at 0°C in the larger

37 37

40 I

region

Y

0 oc

37%+s4

37oc+s4

-

Cl01

RNA

C204

RNA

Figure 8. Gel autoradiographs of V, nuclease digestion data in the hairpin loop region of a mRNA. Cl01 or C204 RNA were partially digested with V, nucleate under the conditions indicated. The portion of each gel displayed extends from A38 (bottom) to A46 (top).

a mRNA Structure fragments (Table l), the TZ nuclease digestion pattern is virtually unchanged, and even more nucleotides appear susceptible to both nucleases (e.g. G43 and A44). There are two possible explanations for why some nucleotides in the hairpin region are sensitive to both single and double-strand-specific enzymes. One are not is that VI and T, nuclease activities absolutely correlated with Watson-Crick basepairing. V, is known to hydrolyze some singlestranded polynucleotides at rates comparable to those for double-stranded RNA (Lowman &, Draper, 1986), and cleavage occurs at some nucleotides in tRNA and rRNA that are involved in tertiary, but not secondary, structure (Auron et al., 1982; Kean & Draper, 1985). Cutting past the ends of helices into single-stranded loops is frequently observed; in a 23-base hairpin, weak V, cleavage is detectable at three of the four loop nucleotides (S. White, unpublished results). T2 and other singlestrand-specific nucleases do cleave rarely within known helices in 16 S rRNA, particularly if the pairing non-Watson-Crick contains helix (Douthewaite et a,Z., 1983; Kean & Draper, 1985). Therefore, unusual secondary and tertiary structure in the hairpin region could be responsible for the nuclease digestion patterns. Another possible reason for overlap between V, and T, nuclease cutting sites is the presence of alternate structures at equilibrium. Many of the nucleotides in the hairpin region are helical in one of the structures shown in Figure 7(a) or (b) and single-stranded in the other. The pseudo-knot structures shown in Figure 7(c) also suggest the possibility of alternate conformations. It is difficult to deduce from structure mapping data alone that a single RNA structure is being observed, but we can make an argument that a unique RNA structure in the hairpin region is trapped by S4 binding (see below). (f) Influence of S4 binding on the structure To determine which secondary features in the a mRNA might be interacting with S4, the nuclease digestion experiments were repeated in the presence of 0.6 PM-s4 (~90% saturation with C204 RNA, x 80% with Cl01 RNA). Distinctive changes are seen in the V, digestion pattern in two regions of the molecule, and are shown in Figures 1 and 8. Small changes in T, nuclease reactivity were seen in the hairpin region with S4 present, but were not observed consistently. We note that the specific activity of T, RNase at pH 7.6 is very low, and the concentrations required to obtain a partial digest were of the order of 0.1 pM. Thus, it is possible that T, RNase affects the binding of S4 to the RNA fragments. A set of five V, cuts in the weak helix region (U6 to UIO) is essentially eliminated by S4 binding (compare the digestions at 37°C with and without S4 in Fig. 1). This could be interpreted either as a direct interaction of S4 with this sequence, or as a melting of the weak helix region. The affinit’y of S4

329

for different a mRNA fragments (Deckman et aE., the latter possibility 1987) supports (see Discussion). S4 also induces distinctive changes in the V, digestion pattern in the same set of hairpin region nucleotides affected by temperature (see Table 1). The pattern becomes more like the digestion of Cl01 RNA at O”C, in that cutting atI U42 becomes very intense, and G43 to A45 become sensitive. The data imply that the entire sequence from A38 to A45 is approximately helical in conformation. About the same cutting pattern is found in either Cl01 or C204 RNA, at either 0°C or 37°C. (g) The in$uence of magnesium ions on a RNA structure In the accompanying paper (Deckham et al., 1987) we found that a mRNA renatured in the presence or absence of Mg2+ has different affinities for S4. To see if Mg 2+ has any specific effect on the mRNA &ructure, we compared V, and T, nuclease digestions at 0°C of the C204 RNA renatured with or without 20 mM-MgCl, present. No specific differences in cutting rates could be detected (data not shown), consistent with our conclusion that the Mg 2+ effect on 54 binding is largely non-specific.

4. Discussion (a) Summary of the a mRNA and a mRNA-S4 structures A model for the c1mRNA secondary structure interacting with S4 is shown in Figure 9; also indicated in the Figure are the approximate free energies of interaction S4 derives from each region, as deduced from the binding studies reported in the preceding paper (Deckman et al., 1987). It should be emphasized that a sequence may contribute to S4 binding either directly by contacting S4, or indirectly by affecting the structure of RNA recognized by S4 elsewhere in the molecule. Here we discuss the structure of each of the three RNA sections in relation to the S4 binding results. (1) The main helix (C23 to U31 paired with A61 to G68) is very stable at 37°C and its structure (as detected by V, nuclease) is unaffected by the presence of S4. The helix is probably extended at one end to include at least three more base-pairs (whether U21 pairs with A72 or A74 is not certain; see Fig. 4). Binding studies with different-sized a mRNA fragments described in the accompanying paper (Deckman et al., 1987) show that the 54 binding site is essentially contained within the region defined by the main helix; the fragment’ C23 to U69 binds with z 9 kcal of the z 9.9 kcal of free energy seen with the intact a mRNA leader. The binding of deletions extending from the 5’ end into the main helix are consistent with the proposed structure. Removal of G22 to U26 reduces the S4 afinity by about 0.7 kcal; these deletions shorten the main helix but the remaining five base-pairs should still be stable. However. a 5’ deletion

UUUUGU

AAUAGUAGGAGUGCAUAGUG

Figure 9. Model for the CLmRNA secondary structure when complexed with S4. Free energy contributions of different sections of the RNA sequence to S4 binding are indicated.

extending through U33 completely eliminates the main helix, and this fragment shows only nonspecific binding. The main helix is therefore essential for S4 recognition, either because of direct interactions with S4, or because the helix stabilizes folding in the hairpin region required for binding. (2) Weak base-pairing takes place between the sequences 5’ and 3’ to the main helix, and probably extends up to the Shine-Dalgarno sequence. The pairing is detectable at 37°C in rather high salt buffer (20 m&r-MgCl,, 350mM-KCl), but must be melted some fraction of the time. Under physiological conditions (37”C, NNO-2 M-salt; Kao-Huang et al., 1977), the potential base-pairing of the ShineDalgarno sequence (Fig. 4(b)) cannot present much of an impediment to initiating ribosomes. S4 does produce a “footprint” in the V, nuclease cuts in the weak helix region; the apparent protection could actually be the result of S4 melting the weak helical structure. The binding studies reported in the accompanying paper show that the sequences Gl to U19 and A70 to Cl39 contribute weakly and independently to S4 binding. If S4 recognized a helix formed from these sequences, disrupting the helix by removing either Gl to U19 or A70 to Cl01 should decrease the binding constant by the same amount. The fact that the 5’ and 3’ termini have additive effects on the S4 affinity suggests that S4 prefers single-stranded structure in this region. (3) S4 binding has a distinct effect on the structure of the hairpin region, though it is difficult unambiguously to deduce the RNA hydrogen bonding pattern from the structure mapping data. We think it most likely that alternative conformations of a pseudo-knot-type structure exist in the absence of 54, and that one of these structures is recognized and trapped by 54. It is significant that the structure of the S4-bound RNA appears the same at 0°C and 37°C in both Cl01 and C204 RNAs, even though the two RNAs show different structures when S4 is not present (Fig. 8). This suggests that a unique RNA conformation is present in the 54 complex, and not an equilibrium between different structures.

Although the distinct RNA structures we have observed in the presence or absence of S4 are detected by changes in V, nuclease reactivity, the structures probably differ in helical conformation, and not in Watson-Crick base-pairing. In other studies of a small nine base-pair helix, we have found that subtle changes in helix backbone structure alter V, cutting rates by as much as an order of magnitude, without changing WatsonCrick base-pairing (S. A. White & D. E. Draper, unpublished results). Therefore, the changes in V, nuclease reactivity that occur in the hairpin region when S4 binds may reflect small changes in the backbone conformation and base stacking in the region. It is also interesting to notice that the requirement for Mg ’ + for full binding of the C204 RNA to S4 could be explained in terms of high negative charge density in the pseudo-knot being preferentially stabilized by Mg’+. The U69 and Cl01 RNAs do not have the same requirement for Mg2+. and cannot form the full pseudo-knot, structure. Altogether, the structure mapping and S4 binding data fit the idea that the hairpin region contains unusual secondary and tertiary interactions that are capable of two or more distinct conformations, and that S4 traps a unique conformation not usually detectable. Some of the subtle changes in S4 affinity with different length c1mRNA fragments may reflect a perturbation of the equilibrium between different hairpin region conformations, so that more binding free energy is required for S4 to trap the conformation it’ recognizes. The expected diameter for a globular protein the molecular weight of S4 is ~40 A, and the measured radius of gyration is 18( +2) A (1 A = 0.1 rim) (Serdyuk et al., 1980). The main helix and hairpin regions can potentially form a continuous helix of 15 base-pairs, roughly 40 A in length and comparable in size to S4. Bends may occur in the helix at bulge positions to reduce the length of the structure, but S4 could interact along the entire RNA structure without any major deformations.

a mRNA Structure

(b) Comparison of the a mRNA and 16 S rRNA structures All ribosomal proteins known to be translational repressors also bind directly to the 16 S or 23 S rRNA. It is simplest to assume that the repressor proteins use essentially the same binding site to recognize both the rRNA and the mRNA site. mRNA and rRNA do compete for repressor binding in all cases that have been studied (for a review, see Draper, 1987; competition between a mRNA and 16 S rRNA for S4 binding was demonstrated by Deckman & Draper (1985)). More direct evidence has been obtained for the Ll and LlO repressors: mRNA and rRNA fragments protected by either protein have been isolated, and these have some primary and secondary features in common (Draper, 1987). The approximate 16 S rRNA region bound by S4 has been defined by mild nuclease digestion of the S4-16 S rRNA complex complex and analysis of the protected RNA fragments obtained (Mackie & Zimmermann, 1975, 1978; Ungewickell et al.. 1975; Khresmann et al., 1980). In each case, a large complex of RNA fragments from the 5’ third of the 16 S rRNA was obtained; the complexes contained many nicks and were not composed of entirely contiguous fragments. The 5’ third of the 16 S RNA (bases 1 to 555) forms one of three large structural domains defined by long-range secondary structure (Noller, 1984). Since S4 is not large enough to interact with the ent)ire domain, it has been thought that S4 interacts with central secondary and tertiary structures to produce the protection observed. Recently Stern et al. (1986) have carried out footprinting experiments with S4 bound to the 16 S rRNA, and found extensive protection within a small but complex region. Five hairpin helices emerge from a small cent’ral loop; a contiguous sequence ext#ending through two of these helices is extensively prot,ected by S4 (C490 to A510), and is presumed by Stern et al. to be the main site of S4 interaction. Two other helices show less extensive protection. Within t#he heavily protected region we have been unable t,o find any significant homology with the a mRNA at the level of either sequence or secondary structure (i.e. positioning of bulges and hairpin loops along a helix). There are several possible reasons for the lack of obvious homology between the a mRNA and the footprinted region of 16 S RNA. The relatively weak specificity of S4 may derive from interactions with only a few RNA bases: the t’wo RN;As may fold in very different tertiary structures that still present t,he same spatial arrangement of recognition features; or t,he footprint may not detect all the S4RNA interactions taking place. All of these factors are probably important. Preliminary results with a series of 16 S rRNA fragments indicate that, if the region Stern d al. (1986) found to be extensively protected by S4 is deleted, specific binding is weakened but not eliminated: specificity apparently derives from three “subdomains” within the first,

331

526 bases of the RNA (J. Vartikar, unpublished results). We therefore think it likely that there are several weak homologies spaced out over a large region of rRNA, and that these homologies may not be obvious in a comparison of the two RNA secondary structures. (c) How does S4 binding injturnce translation? Initiation of translation is known to require a Shine-Dalgarno sequence and an initiation codon with the correct spacing in between (Gold et al., 1981). Discussions of translational repression mechanisms are therefore usually in terms of proteins competing for ribosome binding at the Shine-Dalgarno sequence or initiation codon. or stabilizing an mRNA structure containing these features. The phage T4 regA protein, for instance, is a translational repressor of many T4 genes and probably recognizes a specific sequence overlapping the initiation codon (Karam et al., 1981; Miller et al., 1985). The RI 7 coat protein regulates the translation of the replicase gene by binding to a 21 base stem-loop structure just upst,rea,m from the initiation codon and containing the ShineeDalgarno sequence (Carey et al.. 1983). S4 differs from these other translational ‘repressors in that its strong interaction with t,he a mRNA is upstream (in linear terms) from the ribosome binding site. However. a significant feature of the -secondary structure shown in Figure 9 is the pseudo-knot structure, which physically links the main S4 binding region t)o the ribosome binding sit,e. The structure mapping data also indicate that both S4 and sequences nea,r the initiation codon affect the conformation of the same bases in the hairpin region. This implies that, S4 binding must affect the structure of the initiation codon region. More work is needed to determine the translational repression mechanism, but these observations suggest a way in which S4 binding and ribosome binding may be coupled, without the need to postulate any new features of t,ranslat,ional init’iation. An alternative to direct competition between a translational repressor and ribosomes for binding mRNA is an “entrapment” model in which the repressor allows the ribosome to find the initiation site, but blocks some subsequent step such as tRNA binding or elongation (Draper, 1987). The structure shown in Figure 9 suggests that S4 would be very close to the ribosome in a ribosome-mRNA-S4 complex, and have ample opportunity to disrupt t’ranslation. In the initiation complex, the mRNA is bound to the 3’ terminus of 16 S rRNA, two tRNAs are bound to the mRNA a short distance from the Shine-Dalgarno sequence, and one tRNA is interacting via its anticodon loop with the I6 S rRNA at Cl400 (Prince et al., 1982). Potentially, S4 could interact with these critical sites to block the proper alignment of the RNAs, or prevent dissociation of tRNA or rRNA from the mRNA in subsequent

steps. Determination of the 84-mRPI;A atlinit,y in the presence of ribosomes should be able to distinguish “entrapment” from direct the competit,ion model suggested above.

There is a more extreme case in which mRNA sequences far upstream from the ribosome initiation site affect translation. A set of mutants decreasing translational

efficiency

in the rif operon

is located

over 100 bases upstream from the ribosome binding site (Fiil et al., 1980). This operon is autoregulated by ribosomal protein LlO, and the binding sit,e for this protein is also located about 100 bases upstream from the mRNA initiation site (Friesen et al., 1983; Johnsen et al., 1982: Christensen et al., 1984). An mRNA secondary structure “swikh” coupling LlO binding to t,he formation of helices sequestering the ribosome initiation site has been proposed (Christensen et al., 1984). The LlO and S4 repression systems both suggest that the mRNA plays an active role in mediating translational repression. The possibility of functional RNA “switches” has been discussed for some time. Evidence for switches has been found in different rRNAs, though their functional significance is difficult to establish (Weidner et al.. 1977; Kao & Crothers, 1980; Brimacombe et al., 1983; Christensen et aZ., 1985). Ribosomal proteins are potential triggers for conformational switches, but so far studies of protein-rRNA interactions have found very little perturbation of the RNA structure by proteins (Pl’oller, 1984). Further studies of u mRNA sequence variants may show whether the conformational change induced by S4 binding is related to translational repression. This work was supported by NIH grant GM29048, and D.E.D. acknowledges the support of Research Career Development Award CA01081.

Douthewaite. S.. Christens~~n, A. & Ga.rrett. It. ;\. (19%:~). 169. 249m279. Draper, I>. E. (1987). In Trar&ation,al Regulation of (I’VILP Bq:pression, (Ilan, ,J.. ed.). Plenum Publishing. ?u‘t~~ York. in the press. Draper. D. E.. White, S. A. & Kean. J. til. (1987). Methods h’nzymol. In the press. Ehresmann, C., Stiegler, P., Carbon, C.. ITngewickell. E. & Garrett. R. A. (1980). Eur. J. Biochenb. 103, 339466. Fiil, N. P., Friesen, J. D., Downing, W. 1,. & Dennis. P. P. (1980). Cell, 19, 837-844. Friesen. J. I).. Tropak, M. & An, G. (I 983). (‘rll, 32, 361369. Garrett. R. A. & Oleson. S. 0. (1982). Biochemistry, 21, 48234830. Gold, L.. Pribnow, D., Schneider, T., Shinedling, S., Singer, S. & Stormo, G. (1981). Annu. Rev. Microbial. 35. 365-403. Hertzberg. R. P. & Dervan, P. B. (1982). J. Amer. (Ihem. sac. 104, 313-315. Johnsen, M., Christensen, ‘I’., Dennis. P. P. & Fiil, N. 1’. (1982). EMBO J. 1, 999-1004. Kao, T. H. & Crothers, D. M. (1980). Proc. Nat. Acad. Sri.,

I.:.S.A.

77. 3360-3364.

Kao-Huang, Y., Revzin, A., Butler, A. I’., O’Conner, P., Noble. D. W. & von Hippel, P. H. (1977). Proc. AM. Acad. Ski., U.S.A.

74, 4228-4232.

Karam, ,J.. Gold, L., Singer, B. 6. & Dawson, M. (1981). Proc. Nat. Acad. Sci., G.S.A.

78, 4669-4673.

Kean, J. M. & Draper, D. E. (1985). Biochrmi.stry, 24. 5052-5061. Kean, J. M., White, S. A. & Draper. I). E. (1985). Biochemistry,

24, 5062-5070.

Lockard. R. E. & Kumar, A. (1981). N&. Acids Res. 9, 5125-5140. Lowman, H. B. & Draper, D. E. (1986). J. Biol. Chem. 261, 539C5403. Mackie, G. A. & Zimmermann, R. A. (1975). J. Bio2. Chem. 250, 41OG4112. Mackie, (:. A. & Zimmermann, R. A. (1978). J. Mol. Biol. 121. 17 39. Miller, E. S., Winter, R. B.. Campbell, K. M.. Power. S. I). & Gold. I,. (1985). J. Biol. Chem. 260. 13053G 13059.

References Auron,

P.

E.,

Weber, L. D. & Rich, A. (1982). 21, 4700-4706. Brimacombe, R., Maly, P. & Zwieb, C. (1983). Prog. &cl. Acids Res. Mol. Biol. 28, l-48. Brown, T., Hunter, W. N., Kneale, G. & Kennard, 0. (1986). Proc. Nat. Acad. Sci., U.S.A. 83, 2402-2406. Carey, J., Cameron, V., de Haseth, P. L. & Uhlenbeck, 0. L. (1983). Biochemistry, 22, 2601-2610. Cerretti, D. P., Dean, D., Davies, G. R., Bedwell, D. M. & Nomura, M. (1983). Nucl. Acids Res. 11, 2599-2616. Christensen, T., Johnsen, M., Fiil, N. P. 6 Friesen. J. D. (1984). EMBO J. 3, 1609-1612. Christensen, A., Mathiesen, M., Peattie, D. k Garrett, R. A. (1985). Biochemistry, 24, 2284-2291. Coleman, J., Inouye, M. & Nakamura, K. (1985). J. Mol. Biol. 181, 139-143. Davies, R. W., Waring, R. B., Ray, J. A., Brown, T. A. & Scazzochio. C. (1982). Nature (London), 300, 719724. Deckman, I. C. & Draper, D. E. (1985). Biochemistry, 24, 7860-7865. Deckman, I. C., Draper, D. E. & Thomas, M. S. (1987). J. Mol. Biol. 196, 313-322. Biochemistry,

Edited

Noller. H. F. (1984). Annu. Rev. Biochem. 53, 119--162. Nussinov, R. & Jacobson, A. B. (1980). Proc. Nat. Acad. Sci., U.S.,4. 77, 6309-6313. Pleij, C. W. A., Rietveld, K. & Bosch, I,. (1985). Nucl. Acids Res. 13, 1717-1731. Prince, J. B., Taylor, B. H., Thurlow, D. I,., Ofengand, ,I. & Zimmermann, R. A. (1982). Proc. Nat. Acad. Sci., c:.S.A.

79, 5450.-5454.

Serdyuk. I. N., Gogia, Z. V., Venyaminov, S. Yu., Khechinashvili, N. N., Bushev, V. N. & Spirin, A. S. (1980). J. Mol. Biol. 137, 93-107. Stern. S., Wilson, R. C. & Noller, H. F. (1986). J. Mol. Biol. 192, 101-110. Tinoco, I., Borer, I’. N., Dengler, B., Levine: M. I).. Uhlenbeck, 0. C., Crothers, D. M. & Gralla, .J. (1973). Nature New Biol. 244, 4041. Uchida, T. & Egami, F. (1967). J. Biochem. 61, 44-53. Ungewickell, E., Garrett, R. A., Ehresmann, C., Stiegler, P. & Fellner, P. (1975). Eur. J. Biochem. 51, 165-180. van Stolk, B. *J. & Noller, H. F. (1984). J. Mol. Riol. 180, 151-177. Weidner. H., Yuan, R. & Crothers, D. M. (1977). Nature (London), 266, 193-194. Wurst, R. M., Vournakis, J. N. & Maxam, A. M. (1978). Biochemistry, 17, 4493-4499.

by P. van Hippel