Secondary structure of the 3′ UTR of bicoid mRNA

Biochimie 86 (2004) 91–104 www.elsevier.com/locate/biochi

Secondary structure of the 3′ UTR of bicoid mRNA Christine Brunel *, Chantal Ehresmann UPR 9002 du CNRS, Institut de Biologie Moléculaire et Cellulaire, 15, rue Descartes, 67084 Strasbourg cedex, France Received 31 March 2003; accepted 5 January 2004

Abstract Formation of the Bicoid morphogen gradient in early Drosophila embryos requires the pre-localization of bicoid mRNA to the anterior pole of the egg. The program of bcd mRNA localization involves multiples steps and proceeds from oogenesis until early embryogenesis. This process requires cis-elements in the 3′ UTR of bcd mRNA and successive and/or concomitant critical protein interactions. Furthermore, numerous RNA elements and binding proteins contribute to regulate bcd expression. In the present paper, we investigated the secondary structure of the full length 3′ UTR of the bcd mRNA, using a variety of chemical and enzymatic structural probes. This RNA probing analysis allowed us to give a detailed description of the 3′ UTR of the bcd mRNA and its organization into five well-defined and independent domains (I–V). One prominent result that emerges from our data is the unexpected high degree of flexibility of the different domains relative to each others. This plasticity relies upon the open conformation of the central hinge region interconnecting domains II, III, and IV + V. Otherwise, dimerization of the 3′ UTR, which participates to anchoring bcd mRNA at the anterior pole of the embryo, only results in discrete and local change in domain III. Domain I that contains sites for trans-acting factors exhibiting single stranded RNA binding specificity is mainly unstructured. By contrast, each core domains (II–V) is highly organized and folds into helices interrupted by bulges and interior loops and closed by very exposed apical loops. These elements mostly built specific determinants for trans-acting factors. Besides, these findings provide a valuable database for structure/function studies. © 2004 Elsevier SAS. All rights reserved. Keywords: bcd RNA 3′ UTR; RNA conformation; Probing; Dimerization

1. Introduction The importance of the bcd mRNA 3′ UTR as a repository for signals determining bcd mRNA localization process, polyadenylation, translational regulation and time-triggered degradation has become apparent through abundant recent studies (reviewed in [1,2]). The maternal bicoid gene plays a crucial role in the early development of Drosophila melanogaster. bcd mRNA is transcribed in nurse cells and transported to the oocyte during early stages of oocyte differentiation. Then, it follows a multi-step localization process until its anchoring at the anterior pole of the embryo. The translation of bcd mRNA is kept silenced until the anchoring step is activated, then producing a steep concentration gradient of Bicoid that establishes head and thoracic development (re* Corresponding author. Tel.: +33-3-88-41-70-40; fax: +33-3-88-60-22-18. E-mail address: [email protected] (C. Brunel). © 2004 Elsevier SAS. All rights reserved. doi:10.1016/j.biochi.2004.01.002

viewed in [3]). bcd mRNA is transcribed in nurse cells and transported to the oocyte during early stages of oocyte differentiation. The 3′ UTR encompasses around 800 nucleotides and is organized into domains (I–V) that contain functional elements. Domains IV and V together can direct the earliest phase of transport, and are sufficient for normal localization until embryogenesis. Distinct determinants were identified to be important for the Exuperentia dependant step that governs localization of bcd mRNA at the anterior margin of the oocyte from stage 6 [4,5]. In the latest stages, dimerization through intermolecular base pairing between two complementary loops of domain III, the hairpin loop IIIb and the interior loop IIIa, plays an essential role in the localization process in the embryo [6,7]. The intermolecular interactions, referred as ‘hand-by-arm’ interactions (by opposition to ‘kissing’ loop interactions) were demonstrated by standard disruption/restoration mutation experiments [6,7]. Dimerization was shown to be driven by a two-step mechanism, involving initiation and stabilization [7]. Recently, we

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showed that dimerization is initiated by a more limited number of intermolecular base pairs than expected, leading to a reversible single ‘hand-by-arm’ interaction (‘open’ dimer) [8]. This reversible open dimer is then converted into an irreversible dimer, involving a double ‘hand-by-arm’ interaction (‘closed dimer’). This stabilization step is probably driven by a kinetically controlled mechanism [8]. In the present paper, we investigated the secondary structure of the full length 3′ UTR of the bcd mRNA using a variety of chemical and enzymatic structural probes. Our data allowed us to give a detailed description of the 3′ UTR of the bcd mRNA and its organization into five well-defined and independent domains. This study was carried out on both monomeric and dimeric species, in order to get information about the structural impact of dimerization. Experimental information was provided on bulged nucleotides, interior loops, possible non-canonical base pairs and branching regions. Our results will be discussed on the light of present knowledge of the literature.

2. Material and methods 2.1. Buffers and plasmid construction Buffer D1 (Hepes 50 mM (pH 7.5), 450 mM KCl, 5 mM MgCl2); Buffer M1 (Hepes 50 mM (pH 7.5), 50 mM KCl, 0.1 mM MgCl2); Buffer D2 (50 mM sodium borate (pH 8), 450 mM KCl, 5 mM MgCl2); Buffer M2 (50 mM sodium borate (pH 8), 50 mM KCl, 0.1 mM MgCl2); Buffer D3 (sodium cacodylate 50 mM (pH 7.5), 450 mM KCl, 5 mM MgCl2); Buffer M3 (sodium cacodylate 50 mM (pH 7.5), 50 mM KCl, 0.1 mM MgCl2). Plasmid 875′ used in this study has been described in [7]. 2.2. RNA synthesis and dimerization RNA 875′ was synthesized by T7 RNA polymerase according to [7]. This RNA contains the 875 nucleotides of the bcd mRNA 3′ UTR immediately downstream of the stop codon and 72 additional nucleotides from the vector at the 5′ extremity. It was previously shown that the additional nucle-

A RNase T1

RNase T2

RNase V1

0 1 2 3 0' 1' 2' 3' 0 1 2 3 0' 1' 2' 3' u a c g 0 2 3 0' 1' 2' 3'

-G40 -U50 -G60 -A70 -A80

-U90

-G100

Fig. 1. Enzymatic probing on bcd 3′ UTR. (A) Part of domain I. (B) Part of domain III with loops LIIIa and LIIIb indicated. (C) Central hinge C (cHiC) and part of helix HIVa. Hydrolysis with the indicated RNases was for 2 min (lane 1 and 1′), 4 min (lane 2 and 2′) and 8 min (lane 3 and 3′); incubation control (0 and 0′). Hydrolysis was conducted in buffer M1 (lanes 0–3) or in buffer D1 (lanes 0′–3′). Sequencing lanes (a, u, c, g) were run in parallel.

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otides did not change the dimerization properties of this RNA [7]. RNA 875′ (750 ng) was heated for 3 min at 85 °C in the presence of total tRNA (1.5 µg) in RNase-free water and slowly cooled down to 60 °C. After addition of the fivefold concentrated appropriate buffer, the samples were incubated 30 min at 25 °C prior to be subjected to enzymatic cleavage or chemical modifications. Incubation in buffers D1, D2, D3 yielded 50% of dimeric RNA, while 100% monomers were obtained in buffers M1, M2, M3. 2.3. Enzymatic and chemical probing Enzymatic hydrolyses: incubation was for 2, 4 and 8 min at 25 °C, in the presence of 0.2 U of RNase T1, 0.02 U of RNase T2; 0.2 or 0.05 U of RNase V1, in buffers D1 or M1. Reactions were stopped by phenol/chloroform extraction, followed by ethanol precipitation. CMCT modification: reac-

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tion was for 1, 2 and 4 min, at 25 °C, in the presence of 1.7 µl of CMCT (140 mg/ml), in buffers D2 or M2. Reaction was stopped by addition of 12.5 µM of acetic acid. DMS modification: reaction was for 1.2 and 4 min, at 25 °C, in the presence of 0.5 µl of DMS in buffer D3 or M3. Reaction was stopped by addition of 12.5 µM of b-mercaptoethanol. Alternatively, RNA was treated with DMS and CMCT in buffer D3 and D2, respectively, and the modified monomeric and dimeric RNA species were fractionated by electrophoresis on native 1% agarose gel at room temperature. Electrophoresis buffer and gels contained 45 mM Tris borate (pH 8.3) and 0.05 mM MgCl2. Monomers and dimers were purified from the gel slices by phenol extraction and ethanol precipitation in the presence of glycogen as a carrier. This procedure allowed us to analyze pure dimeric and monomeric species under identical ionic conditions.

C

B RNase T1 0 1 2 3 0' 1' 2' 3'

RNase T1

u a c g

0 1 2 3 0' 1' 2' 3' u

RNase V1 c g 0 1 2 3 0' 1' 2' 3'

-A250 -G260

LIIIb

-U270

G360-

U370-

C380-

-C280 U390-

-U290

-U300

LIIIa

-G310

-3C20

Fig. 1 (continued)

cHiB

C350-

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A

B DMS

CMCT

A120- 0' 1' 3' 0' 1' 3' u a c g 0 1 2 3 0' 1' 2' 3'

cHiA

A130C140-

U150-

A C U C C A G U U A A C U C U A U

DMS Monomer Dimer 0' 1' 2' 3' 0' 1' 2' 3' u a c g

-G200 -A210 -A220 -U230

G160-

-U240

C170-

C G200A210A220U230-

CMCT u a c g 0 1 2 3 0' 1' 2' 3'

A C U C C A G U U A A C U C U A U

cHiA

G180-

U240-

Fig. 2. Chemical probing on bcd 3′ UTR. (A) Part of domain 1. (B) and (C) Central hinge A (cHiA) and part of helix HIIIb. Modification with DMS and CMCT was for 1 min (lane 1 and 1′), 2 min (lane 2 and 2′) and 4 min (lane 3 and 3′); Incubation control (0 and 0′). CMCT modification was conducted in buffer M2 (lanes 0–3) or D2 (lanes 0′–3′). In the case of DMS, RNA was modified in buffer D3 and monomeric and dimeric RNA species were purified and analyzed (see Section 2). Sequencing lanes (a, u, c, g) were run in parallel.

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A U290-

B

DMS Monomer Dimer u a c g

95

CMCT

0' 1' 2' 3' 0' 2' 3'

a g c u 0 1 2 3 0' 1' 2' 3'

U290-

*

U300-

U300-

* AC

G310-

LIIIa

U U U G G G C U U A

C320-

C320U330-

LIIIa

C A U U U G G G C U U A

G310-

U330-

A340A340U A U C G U C C C A C A U U U C

C

cHiB

C350-

D

DMS Monomer Dimer u a c g 0' 1' 2' 3' 0' 2' 3'

U270-

*

U270-

C280-

A C U A A A G C C C G

LIIIb

U290-

*

A C U A A A G C C C G

LIIIb

C280-

CMCT u a c g 0 1 2 3 0' 1' 2' 3'

U290-

Fig. 3. Chemical probing on bcd 3′ UTR. (A) Part of domain III (with loop LIIIa) and central hinge B. (B) Part of domain III with loop LIIIa; (C) and (D) Part of domain III with loop LIIIb. Modification with DMS and CMCT was as in Fig. 2. The asterisks denoted Gs modified at position N7 with DMS revealed after cleavage with sodium borohydrure [11].

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2.4. Detection of modifications and cleavages Modified bases or chain scissions were detected by primer extension with avian myeloblastosis virus reverse transcriptase as described previously [9] using various 5′-end [32P]-labeled primers complementary to residues 134–150, 194–200, 284–300, 374–390, 464–480, 554–570, 644–660, 764–780, 824–840. Analysis of the generated cDNA fragments were as described in [10]. For each reaction, a control was treated in parallel, omitting the chemical or enzymatic reagent. Dideoxynucleotide sequencing reactions were run in parallel to allow identification of the modified positions.

3. Results 3.1. Experimental strategy: chemical and enzymatic probing and derivation of the secondary structure model The conformation of RNA 875′ was probed using a variety of enzymatic and chemical probes (for details, see [11,12]). RNase T1 was used to map unpaired guanines, RNase T2 unpaired adenines and to a lesser extend unpaired uridines, and RNase V1 double-stranded or stacked regions. Watson– Crick positions were tested with DMS (A(N1) and C(N3)), and CMCT (G(N1) and U(N3)). The effects induced by

intermolecular RNA–RNA interaction were tested under two salt conditions, either preventing or favoring the RNA–RNA association. Monomeric species (95–100%) were obtained in low-salt buffers (containing 50 mM KCl and 0.1 mM Mg2+). A 50% mixture of dimeric and monomeric species was obtained in high-salt buffers (containing 450 mM KCl and 5 mM Mg2+). The resulting dimers correspond to the stabilized forms (the stable ‘closed’ dimers) and do not dissociate after dilution for hours [7,8]. An alternative procedure was used to analyze purified dimers and monomers under the same ionic conditions. Chemical modification was done on the half to half mixture of monomers and dimers under high-salt conditions. Then, the modified monomers or dimers were fractionated by gel electrophoresis under native conditions before analysis. The complete molecule was analyzed by this procedure with DMS, and domain III with CMCT. Chemical and enzymatic probing were repeated two to four times in independent experiments. Representative gel autoradiographs are shown in Figs. 1–5, and results are compiled in Fig. 6 and Table 1. Experimental data were used to derive a secondary structure model of the bcd 3′ UTR RNA. The construction of the secondary structure model was assisted by computer prediction allowing us to generate optimal and suboptimal folds [13,14]. The secondary structure motifs consistent with reactivity data were selected, and the folding program was run

CMCT A340C350G360-

U370-

C380-

a g c u 0 2 3 0' 1' 2' 3'

U A U C G U C C C A C A U U U C G G A A A U U A

cHiB

A

U390-

Fig. 4. Chemical probing on bcd 3′ UTR. (A) Central hinge B (cHiB) and part of helix HIVa. (B) and (C) Part of domain V. Modification with DMS and CMCT was as in Fig. 2.

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again, forcing the selected secondary structure elements to be integrated. The bcd mRNA 3′ UTR sequences from nine Drosophila species and from Musca domestica were compared (Fig. 7). Remarkably, all the bcd 3′ UTR RNA analyzed in this study share a common global secondary structure, in agreement with previous analyses using a smaller set of sequences [15,16]. However, sequence divergence was poorly informative in term of covariation. In particular, the length of helices and loops, the bulge positions within helical regions are not conserved through the different species. Nevertheless, this

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analysis underscored the presence of several stretches of conserved nucleotides through the Drosophila genus (Fig. 7). Functional relevancy of these sequences will be discussed below. 3.2. Secondary structure model of the monomer Enzymatic and chemical probing experiments were conducted on monomeric RNA obtained in low-salt buffers. Notably, a similar DMS modification profile was observed when RNA was treated in high-salt buffer before isolation of

C

B

DMS Monomer

CMCT u a c g

Dimer

0' 1' 2' 3' 0' 1' 2' 3' u a c g

0 1 2 3 0' 1' 2' 3'

A490-

-A490

A500-

-A500

G510-

-G510

U520-

-U520

U530-

-U530

-C540

C540-

Fig. 4 (continued)

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A DMS Monomer Dimer

CMCT

0' 1' 2' 3' 0' 1' 2' 3' u a c g

0 1 2 3 0' 1' 2' 3'

C620C630G640-

A660-

A670-

U680-

A690-

A C U A U C A U A G C U C A C A U U C U A U U U A

cHiC

U650-

A700Fig. 5. Chemical probing on bcd 3′ UTR. (A) Central hinge C (cHiC) and parts of flanking helices HIVa and HII. Part of domain IV. Modification with DMS and CMCT was as in Fig. 2.

monomers. There is a satisfying agreement between reactivity data and the derived secondary structure model (Fig. 6). As a general rule, nucleotides in helices were unreactive to chemicals, but low or moderate reactivity was sometimes observed, essentially in A–U pairs, reflecting some breathing of helices portions of lower stability or at the extremities of helices. It should also be noticed that A is more reactive to DMS than C, and U more reactive than G to CMCT [11]. If loops generally displayed a high degree of reactivity to chemicals, all nucleotides were rarely evenly reactive. The presence of weakly reactive or unreactive nucleotides reflects a certain degree of organization, such as non-canonical interactions or stacking of nucleotides (i.e. in the helical continuity of flanking helices). Otherwise, the reactivity of nucleotides in loops or in single stranded connecting regions was most often associated to the occurrence of RNase T1 and/or T2 cuts. RNase T1 cuts were only rarely found in helical parts. In this case, they reflect local breathing or the occurrence of secondary cleavages. These cleavages are usually

weak as compared to those observed in single stranded regions (e.g. see cuts at G284, G285 and G288 in Fig. 1B). RNase V1 cuts were essentially found in helical regions or at the boundaries of loops. In some cases, RNase V1 cuts were found in apparently single-stranded regions (e.g. domain I, region connecting domains III and IVa, and at position 504 in the large asymmetrical loop in domain V). Such cleavages are generally interpreted to reflect conformational heterogeneity or stacking of unpaired nucleotides [11,12]. However, the reactivity of a number of nucleotides could not be determined due to the presence of stops or pauses of reverse transcriptase that are generally assigned to cleavages or to stable structural elements. Our data mostly validate the existence of the five independent domains named I–V previously proposed on the basis of computer prediction and sequence comparison [15,16]. The structured core is constrained by domain II, which pairs distant nucleotides (172–204 and 722–690). Domains III, IV (divided into two subdomains IVa and IVb) and V fold into

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B CMCT 0 1 2 3 0' 1' 2' 3'

u a c g

-G600

-U610

-C620

-C630

-G640 Fig. 5 (continued)

complex stem–loop structures, with their helical regions interrupted by bulges and interior loops of unequal size. Continuous helices rarely exceed a full turn (11 base pairs). However, important divergence was found in the central hinge region connecting domains II, III and IV/V, where the previously proposed model predicted a more compact structure (see insert of Fig. 6). Two stem–loop structures were proposed in the region connecting domains III and IV on one side, and IV and II on the other side. The first one involved a regular helix (356– 362/369–375) closed by a six-base loop (363–368), and the second one was closed by an irregular helix (654–667/671–

687) interrupted by an interior loop (see insert of Fig. 6). However, the general high level of reactivity of nucleotides UUCGGAA362 (Figs. 4A and 6) and AUUCUAUUU687 (Figs. 5A and 6) forming the respective 3′ and 5′ strands of the two putative hairpins did not support their existence. On the other hand, the low reactivity of the complementary sequences UAUUCCG373 (Figs. 4A and 6) and CGGAAUA663 (Figs. 5A and 6), and the presence of RNase V1 cuts in these sequences (Figs. 1C and 6), were in favor of the existence of an alternative helix extending helix HIVa. This stem is separated from helix HIVa by an interior loop encompassing nucleotides 372–375 and 652–656, which is sup-

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Fig. 6. Secondary structure model of the central part of bcd mRNA 3′ UTR. The nomenclature of domains is that previously used [6,7,15,16] and cHiA, cHiB and cHiC stand for central hinge A, B, C, respectively. Enzymatic cleavage by RNase T1 is indicated as an arrow ended by a square, RNase T2 as an arrow ended by a circle, RNase V1 as a simple arrow. The intensity of cleavage increased with size and darkness of the arrows. Chemical modification on Watson–Crick positions are noted as highly reactive (red full circles) moderately reactive (orange full circles) marginally reactive (yellow full circles) not reactive (blue full squares). Non-determined nucleotides are not encircled. The structure in the yellow insert shows the secondary structure model of the central hinge region previously predicted [15,16]. Nucleotides involved in a divergent secondary structure in our experimental derived model are shown in red.

ported by the reactivity of nucleotides CCAAA656 (Figs. 5A and 6). Otherwise, a few base pairs were proposed between nucleotides flanking helix HIIIa (GU212/AC353, AC216/GU348, UA220/UA344), resulting in a very irregular helical region (see insert of Fig. 6). Only a few of the concerned nucleotides were unreactive (C216, C218) (Figs. 2B and 6), suggesting that these base pairs either do not exist or form unstable interactions. The left-unpaired nucleotides forming the central hinge (denoted cHiA, cHiB and cHiC in Fig. 6) were essentially reactive to chemicals (Figs. 2B, C, 3A and 4–6), with a few RNase T1 and T2 cuts (at A210, G211, G359 and G360). Notably, several RNase V1 cuts were also observed, overlapping two strong RNase T1 cuts at GG360 (Fig. 1C). They might result from a stacked configuration of the concerned purine-rich sequence or to the

presence of a possible alternative metastable interaction, involving base pairing between UCC350 and GGA361. In conclusion, our results indicate that the central hinge is essentially unstructured and flexible, resulting in a high degree of freedom of domains II, III and IV + V relative to each other. By contrast, the three-way junction between helices HIVa, HIVb and HVa (small hinge) is more compactly organized, indicating that domains IV and V form a structural entity. In contrast to nucleotides 172–722, which fold into a highly structured core, the flanking 5′ and 3′ terminal regions (nucleotides 1–171 and 723–760) forming domain I, are mostly single-stranded. However, RNase V1 and singlestranded specific modifications or cleavages were concomitantly observed in several distinct areas (Fig. 1A and

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Table 1 Reactivity of domain I nucleotides. Four level of reactivities toward DMS and CMCT or accessibilities toward enzymatic probes are estimated: (+++) highly reactive, (++) moderately reactive, (+) marginally reactive, (o) unreactive. An absence of symbol indicated an absence of information. White lettered nucleotides in dark boxes indicated NRE elements

Table 1), suggesting the occurrence of alternative metastable conformations. 3.3. Dimerization does not induce important conformational rearrangement The RNA was then analyzed in high-salt buffers that favor dimer association. Although RNase T1, T2 and V1 hits were the same in low- and high-salt buffers, some intrinsic variability in the level of cleavage was observed among experiments (Figs. 1A and 2B, D). This behavior most likely accounts for (i) the sensitivity of RNase activities to ionic conditions; (ii) the stabilizing effect of monovalent and divalent ions on RNA conformation; (iii) experimental fluctuations. Notably, the most important variability was observed with RNase V1, which requires Mg for catalysis, and to a less extent with RNase T2. RNase T1 appeared to be less sensitive to experimental conditions. Despite the observed fluctuations, the cleavage patterns did not reveal important reorganization of the RNA upon dimerization. In line with this view, DMS and CMCT modification yielded very similar patterns in low- and high-salt buffers. We only observed a general CMCT decrease in high-salt buffer (Figs. 2A, C, 3B, D, 4A, B and 5A, B). Thus, it was important to compare DMS modification profiles of pure monomers and dimers fractionated after modification in high-salt buffer, because results were not biased by different experimental conditions. In this case, no significant differences could be observed, confirm-

ing that dimerization did not trigger important conformational rearrangement. However, we failed do detect a clear footprint of the dimer in the dimerization domain, in particular in loops IIIb and IIIa. The only exception concerned RNase T1 cleavage at G317, G318 and G319 in loop IIIa of domain III, which displayed a slight but reproducible reduction upon dimerization (Fig. 1B). On the other hand, the RNase T1 cut at G279 in loop IIIa did no show any detectable decrease, but a possible reduction might have been counterbalanced by its very high reactivity. The most unexpected result was the absence of protection to chemicals in the two complementary sequences AAGCCC282 and GGGCUU322 (Fig. 3), with the only possible exception of G279 (Fig. 3D). However, it should be noted that among the six putatively concerned nucleotides of each sequence, C280 in loop IIIb, and G317, G318 in loop IIIa were already unreactive at Watson–Crick positions in the monomer (Fig. 3). Moreover, no information could be available for C281, C282 and G319, C320 due to the presence of reverse transcriptase pauses (Fig. 3). Beside, A277, A278, located at the 5′ edge of loop IIIb, and U321 and U322 at the 3′ edge of loop IIIa, were reactive in both species (Fig. 3C, D). The fact that the reactivity of UU322 and AA278 was unchanged in the dimer argues against the formation of stable interactions between these nucleotides. These findings suggest either that base pairing is not as extended as expected or that the extremity of the postulated intermolecular helix (UU322/AA278) is breathing.

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Fig. 7. Conserved sequence within the central part of the bcd mRNA 3′ UTR. Sequence conservation through the nine Drosophila species (D. melanogaster (accession number X07870), D. sechellia (M32124), D. simulans (M32123), D. teissieri (M32121), D. virilis (M32122), D. picticornis (M32126), D. heteroneura (M32125), D. pseudoobscura (X55735), D. suboscura (X78058) and M. domestica (AJ297852)) is indicated by red letters (100%), green letters (90%) and black letters (80%). Known cis-acting elements are schematically indicated.

This absence of clear changes in the two autocomplementary loops is partly explained by the fact that RNA 875′ only yielded 50% of dimerization. In addition, interpretation was spoiled by intrinsic limitations of probing experiments (nonreactivity of some nucleotides, pauses of reverse transcription). Nonetheless, recent results indicated that dimerization is initiated by only a limited number of nucleotides, namely CCC282/GGG319 [8]. In particular, footprinting experiments conducted on the isolated domain III (RNA III) revealed a clear protection to DMS modification at CCC282 in loop IIIb, while C320 remained fully reactive in loop IIIa. Unfortunately, no information could be available for these nucleotides in the present experiments. Moreover, we showed that the initial reversible dimer only involves a single hand-to-arm interaction (‘open’ dimer), while stabilization is triggered by the formation of a second hand-to-arm interaction, resulting in a ‘closed’ dimer [8]. This transition occurs in large RNA fragments, while RNA III preferentially forms multimers through hand-to-arm interactions. The evidence that the two interactions simultaneously occurred on the same RNA molecule was demonstrated for the first time by the use of heterolength-dimers between RNA lacking domain I (RNA _I) and a truncated RNA (RNA _(I, IV, V) lacking domain I, domain V and most part of domain IV [8]. The resulting heterodimers formed stable dimers at high

yield (up to 95%) that allowed visualization of unambiguous RNase T1 protection in both loops IIIb and IIIa (see Fig. 7 of [8]). 4. Discussion bcd 3′ UTR contains cis-acting elements involved in a variety of functions: mRNA localization, polyadenylation, translational regulation and time-triggered degradation (reviewed in [1,2]). RNA probing allowed us to get the first experimental insight into the folding that frames bcd 3′ UTR. Our data confirm the partition of the bcd 3′ UTR into welldefined and structured domains. One prominent result is the rather unexpected high degree of flexibility regarding domains orientation, resulting from the absence of stable interactions in the central hinge region (Fig. 6). This conclusion constitutes the major divergence with the in silico predicted folding model [15,16], in which the articulation of the different domains was constrained by extensive base pairing (Fig. 6). This plasticity might represent a pivotal feature for the numerous functions that govern the fate of the mRNA. Indeed, the various elements of the 3′ UTR should participate in successive and/or concomitant critical interactions with different protein factors from the early localization process until controlled expression and final degradation.

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Deduced from our data, domain I is poorly structured. Metastable hairpins may form, accounting for the few RNase V1 cuts in conjunction with the chemical modification pattern, in particular between positions 105 and 130. A parallel behavior occurs in silico as domain I folds into numerous secondary structures exhibiting a very close range variation of free energy (DG). Early experiments demonstrated that domain I is dispensable for the complete localization process, suggesting that all cis-acting sequences were located within the core (domain II–V) [17,18]. However, a more recent work demonstrated that the 5′ part, immediately following the stop codon, contains a cis-element responsible for time-dependent degradation of the bcd mRNA during cellularization of the blastoderm [19] (Fig. 7). This element is able to induce degradation when inserted into heterologous RNAs, independently from the structural context [19]. A bipartite nanos-responsive-element (NRE) sequence is located at nucleotides 51–55 and 61–66 and a downstream B box at positions 80–85 [20] (Fig. 7). These NRE are indeed operational, temporally regulating bcd mRNA expression in a Pumilio-dependent manner [20]. Interestingly, the NRE nucleotides were fully reactive to chemicals at Watson–Crick positions and to single strand-specific RNases (Table 1), consistent with the single stranded RNA binding specificity of Pumilio family proteins. The 3′ part of domain I also contains two putative polyadenylation signals that also need to be accessible to the maturation apparatus [15,16,21]. The structured core of the 3′ UTR mRNA was found to contain important signals for different steps of the bcd mRNA localization during oogenesis and embryogenesis. Remarkably, all these cis-acting elements are located within strictly conserved sequences through the Drosophila genus (Fig. 7). Our study provides new useful information on the secondary structure of these sequences. In domain V, mutagenesis experiments revealed that both the secondary structure (maintaining helical regions) and specific nucleotides (U480, A481, U486, C487) are required for the early steps of transport [5]. According to our data, these nucleotides are located in two small interior loops, most likely available for specific interaction with proteins of interest. Indeed, unlike DNA, regular RNA helices are unsuited to provide specific determinants for protein recognition, unless they contain irregularities, bulged nucleotides or distortion of grooves [22]. In this line, the deleterious effect of the U486 to G mutation would result from the formation of a G486–C447 base pair changing the local structural context. Otherwise, distinct determinants (U460, A481, A492, U495, A496, A498, A504) were identified to be important for the Exuperentia dependant step that governs localization of bcd mRNA at the anterior margin of the oocyte from stage 6 [4,5]. From our data, these nucleotides map in or close to the apical loop closing helix Vb and in the large interior loop between helices HVa and HVb, which are both readily accessible in the naked RNA. Our data clearly indicate that no major rearrangement results from dimerization. Thus, dimerization does not seem

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to interfere with the conformation of other cis-acting signals. Moreover, because of the high flexibility between domains, dimerization is not expected to impede the accessibility of these elements to their different interactors. It might even favor or stabilize interactions by generating an intrinsic duplication of regulatory sites. However, the lack of clear footprint within the complementary sequences of loops IIIa and IIIb, mainly due to internal limitations, prevented to precisely determine the number of nucleotides engaged in intermolecular base pairs. In conclusion, our results provide detailed information on the structural context of the various regulatory elements. They constitute a valuable data bank for guiding the design of site-directed mutagenesis and will be useful for further structure/function investigation.

Acknowledgements We thank P. Romby for helpful comments on the manuscript, C. Wagner, F. Brulé and C. Isel for valuable discussions. We thank Christian Massire and Eric Westhof for help with the COSEQ program. This work was supported by the CNRS.

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