Nucleic Acids Research

Chaveroche, Marie-Kim;Ghigo, Jean-Marc;d’Enfert, Christophe

doi: 10.1093/nar/28.22.e97pmid: 11071951

Abstract The construction of mutant fungal strains is often limited by the poor efficiency of homologous recombination in these organisms. Higher recombination efficiencies can be obtained by increasing the length of homologous DNA flanking the transformation marker, although this is a tedious process when standard molecular biology techniques are used for the construction of gene replacement cassettes. Here, we present a two-step technology which takes advantage of an Escherichia coli strain expressing the phage λ Red(gam, bet, exo) functions and involves (i) the construction in this strain of a recombinant cosmid by in vivo recombination between a cosmid carrying a genomic region of interest and a PCR-generated transformation marker flanked by 50 bp regions of homology with the target DNA and (ii) genetic exchange in the fungus itself between the chromosomal locus and the circular or linearized recombinant cosmid. This strategy enables the rapid establishment of mutant strains carrying gene knock-outs with efficiencies >50%. It should also be appropriate for the construction of fungal strains with gene fusions or promoter replacements. Received July 26, 2000; Revised and Accepted September 29, 2000. INTRODUCTION Gene manipulation in filamentous fungi is limited by the low efficiency of homologous recombination upon DNA-mediated transformation and the high frequency of ectopic integrations of the transforming DNA molecule: a minimum of 1 kb of DNA homologous to the target site is often required to achieve efficiencies of homologous recombination ~10% (1,2). This will become a major limitation for post-genomic studies once whole genome sequences of filamentous fungi become available. It follows that construction of knock-out mutants by gene replacement often requires several subcloning steps prior to transformation of the fungus and is therefore a lengthy process. An alternative strategy has been proposed where knock-out constructs are directly placed on a cosmid carrying a genomic region encompassing the gene of interest, thus providing large regions of homology with the genome. This can be achieved either by in vivo or in vitro transposon mutagenesis (L.Hamer and J.E.Hamer, 20th Fungal Genetics Conference, Asilomar, USA) or by recombination between the cosmid and a PCR product in Saccharomyces cerevisiae (J.A.Sweigard, 20th Fungal Genetics Conference, Asilomar, USA). However, these two strategies, which have so far not been described in detail, have drawbacks since their application would either result in random insertions or require the development of novel genomic libraries based on Escherichia coli–S.cerevisiae shuttle vectors. Recently, it has been shown that expression of the E.coli RecE and RecT proteins or the corresponding Redα(exo) and Redβ(bet) proteins of phage λ together with the Redγ(gam) protein can drastically promote homologous recombination in E.coli. Using this approach it has been possible to achieve allelic exchanges for genes located on the E.coli genome or an episome by recombination with a PCR product containing homology arms of 50 nt (3–6). We have developed a similar strategy where expression of Redα, Redβ and Redγ is used to promote recombination in E.coli between an amplified bi-functional transformation marker and a selected cosmid carrying an Aspergillus nidulans genomic region. The circular or linearized cosmid is then used to transform A.nidulans yielding transformants with the appropriate gene replacement at frequencies up to 60%. This methodology should be applicable to most filamentous fungi and can be used to introduce highly specific gene modifications in fungal genomes. MATERIALS AND METHODS Strains and culture conditions Escherichia coli strain KS272 [F– ΔlacX74 galE galK thi rpsL ΔphoA (PvuII)] was used for propagation of the recombination vector pKOBEG and of A.nidulans cosmids. Conditional R6Kγ origin plasmids were maintained in the pir+ host DH5α λpir [F–endA1 hsdR17 supE44 thi-1 recA1 gyrA relA1 Δ(lacIZYA-argF)U169 deoR (ϕ80dlac(lacZ)M15)(λpyr+)]. Escherichia coli strains were propagated in LB medium or LBLS (1% bacto-tryptone, 0.5% yeast extract, 0.5% NaCl, pH 7.5) medium when selection for zeocin resistance was applied. The β-lactam antibiotic carbenicillin (100 µg/ml), chloramphenicol (25 µg/ml), spectinomycin (100 µg/ml) and zeocin (50 µg/ml; Invitrogen) were added to the growth medium when required. l-Arabinose or d-glucose were added as indicated to modulate expression of genes under control of the pBAD promoter (7). Aspergillus nidulans strain FGSC773 (wA3 pyroA4 pyrG89) was obtained from the Fungal Genetics Stock Centre (University of Kansas, Kansas City, USA). Growth conditions for A.nidulans strains have been described (8). Scoring of the acid trehalase phenotype was achieved by replica plating of FGSC773 transformants on minimal medium containing 1% glucose or 1% trehalose as the carbon source (8). DNA manipulations DNA manipulations were according to Sambrook et al. (9) and Ausubel et al. (10). Transformation of calcium–manganese-treated E.coli was as described (11). Electro-competent E.coli cells were obtained by washing exponentially growing cells three times with ice-cold distilled H2O and concentrating 300-fold with 10% glycerol. Electroporation was carried out in 0.2-cm electroporation chambers and using a BioRad GenePulser II set to the following parameters: 200 Ω, 25 µF and 2.5 kV. Shocked cells were diluted 3-fold in LB, incubated for 1 h at 30°C without aeration and subsequently plated on appropriate media. Plasmid pTP223 (3), pBAD18 (7), pKO7, a derivative of pKO3 (12), pHP45Ω-Spc (13) and pGP704Not, a derivative of pGP704 (14), were obtained from K. Murphy, J. Beckwith, G. Church, H. Krish and D. Mazel, respectively. pUC4K and pEM7-zeo were purchased from Pharmacia and Invitrogen, respectively. pAfpyrG2, pTRE11 and pTRE12 have been described previously (8,15). Construction of the recombination plasmid pKOBEG was as follows. The NsiI–HindIII fragment of pBAD18 carrying the pBAD promotor (7) was subcloned into PstI–HindIII-digested pUC18, yielding pUCaraC-pBAD. The λ Red region of pTP223 was amplified using oligonucleotides EBGNHe-5 (5′-CCCGCTAGCGAAAAGATGTTTCGTGAAGC-3′) and EBGh3 (5′-GGGAAGCTTATTATCGTGAGGATGCGTCA-3′). The resulting 1960 bp PCR product was digested with NheI and HindIII and subcloned into NheI–HindIII-digested pUCaraC-pBAD, yielding pUCaraC-P-EBG. The KpnI–HindIII fragment of pUCaraC-P-EBG was then subcloned into KpnI–HindIII-digested pKO7, yielding pKOBEG. pCDA18 was obtained by subcloning the EcoRI 2.1 kb spectinomycin-resistance gene carried by pHP45Ω-Spc at the EcoRI site located in 3′ of the Aspergillus fumigatuspyrG gene in pAfpyrG2. pCDA19 is a derivative of pCDA18 where the NotI site has been replaced by EcoRV using an adaptor of the following sequence, 5′-GGCCGATATC-3′. pTRE18 was then obtained by inserting the 4 kb EcoRV of pCDA19 into EcoRI–PmlI-digested pTRE11 that had been treated with T4 DNA polymerase. pCDA21 was obtained by subcloning the 1.9 kb EcoRV–NotI fragment of pAfpyrG2 carrying the A.fumigatus pyrG gene and the 0.5 kb NotI–EcoRI fragment of pEM7-zeo carrying a zeocin-resistance gene into EcoRV–EcoRI-digested pGP704Not. Allelic replacement on cosmids Electrocompetent cells of a transformant carrying pKOBEG (CmR) and cosmid W14C08 (AmpR KmR), a derivative of pWE15 which carries an A.nidulans genomic region encompassing the treA gene (8), were prepared from a culture containing 0.2% arabinose in order to induce the expression of the red genes. Electrocompetent cells (100 µl) were electroporated with ~100 ng of the gel-purified 7 kb NotI fragment of pTRE18 or of a dialyzed PCR product obtained by amplification of pCDA21 with the following oligonucleotides: treFzeo (5′-GCTGAAGGTTCCTTTCTTGATCCTCTCTGTGGGTTCAGAAAGCGTTCAAAggaattctcagtcctgctcc-3′) and treBpyr (5′-CTCAGATTCTATGCTAAACTCGCTCATACTACTCTACATGATATTCTACAgaattcgcctcaaacaatgc-3′). These oligonucleotides have 50 bp of homology to the 5′ or 3′ non-coding region of the A.nidulans treA gene (upper case) followed by 20 bp of homology to the zeocin-resistance or A.fumigatus pyrG genes carried by pCDA21 (lower case), respectively. Amplification was performed in a 100 µl reaction containing 200 ng pCDA21, 25 nmol dNTPs, 68 pmol treFzeo, 63 pmol treBpyr, 5 U rTaq (Pharmacia) and using the following protocol: a denaturation step at 93.5°C for 5 min followed by four cycles of the following steps: denaturation at 93°C for 30 s, annealing at 58°C for 2 min, extension at 72°C for 3 min and 24 cycles of the following steps: denaturation at 93°C for 30 s, annealing at 60°C for 2 min, extension at 72°C for 3 min. A last elongation step was done at 72°C for 10 min. Shocked cells were plated on LB medium containing kanamycin and spectinomycin (NotI fragment) or LBLS medium containing kanamycin and zeocin (PCR product) and incubated at 30°C. Transformants were colony-purified once at 42°C on medium containing kanamycin and spectinomycin or zeocin and then tested for chloramphenicol resistance to test loss of pKOBEG. The resulting cosmids were further characterized by restriction enzyme digestion and PCR analysis using two sets of primers located either in the replaced region (tre#5, 5′- AGGACTCGGTCGATCGCT-3′; tre#6, 5′-CCGGATGGGAGGCGACGA-3′) or in the vicinity of the replaced region (tre#1, 5′-ATCCACTTGTTCATCGTC-3′; tre#2, 5′-AGGCAGAAAGATGATCTC-3′). Allelic exchange in A.nidulans Transformation of A.nidulans strain FGSC773 was according to the procedure of Osmani et al. (16) using 5 µg of circular pTRE18, cTRE19 and cTRE20 or cTRE20 DNA that had been digested with SfiI or ClaI or treated with DNase I for various times and in the presence of manganese in order to achieve partial linearization at random sites on the cosmid (10). Uridine/uracil prototrophic transformants were scored for their ability to grow on minimal medium containing trehalose as the sole carbon source. Diffuse growth on this medium was indicative of an allelic replacement at the treA locus (8). Allelic replacement at the treA locus was tested by PCR using primers tre11 (5′-ATTGGTCTTCTGGGATG-3′) and tre10 (5′-CAAAAGCCGTAAAACTTACA-3′) which generate a 434 bp fragment only in strains that have an intact treA gene or by Southern hybridization of EcoRI-digested genomic DNA. In this case, a 2.0 kb EcoRV fragment of pTRE10 (8) corresponding to the 3′-end of treA was used as a probe and detected an 8.0 kb fragment in treA+ strains and a 4.7 kb fragment in strains containing the treAΔ::zeo/pyrG allele integrated either at the treA locus or at ectopic sites. The fact that integration had not occurred at ectopic sites or in multi-copy in treA– strains is confirmed by comparing the intensity of the signals obtained using the treA probe and a probe for a single copy gene. RESULTS Establishment of a recombination system Molecular genetics of the filamentous fungus A.nidulans has been facilitated by the availability of chromosome-specific cosmid libraries and their corresponding physical maps (17,18). These libraries were constructed in the pWE15 and LORIST2 cosmid vectors which carry a ColE1 origin of replication and ampicillin and/or kanamycin resistance genes. Furthermore, DNA-mediated transformation of A.nidulans takes advantage of selective markers (e.g. pyrG which encode orotidine-5′-monophosphate decarboxylase and confers prototrophy when introduced into an A.nidulans pyrG uridine/uracil auxotroph) that are not functional in E.coli. Two plasmids were therefore designed for in vivo modification of cosmids that could be subsequently used to transform A.nidulans (Fig. 1). Plasmid pKOBEG (Fig. 2A) is a thermosensitive replicon that carries the λ phage redγβαoperon (3)expressed under the control of the arabinose-inducible pBAD promotor (7). Because pKOBEG is a derivative of pSC101 and confers chloramphenicol resistance it can be propagated in E.coli together with most ColE1-derived plasmids. The functionality and inducibility of the Red functions encoded by pKOBEG were confirmed by inactivating the dnaK gene on the E.coli genome using a kanamycin-resistance gene flanked by 1000 bp of DNA homologous to the 5′ and 3′ regions of the dnaK gene. While none of the KmR transformants obtained from glucose-grown cells showed the appropriate gene replacement, at least 80% of the transformants obtained from arabinose-grown cells had occurred from a gene replacement at the dnaK locus (data not shown). Plasmid pCDA21 (Fig. 2B) contains a bi-functional transformation marker (zeo/pyrG) constructed using a zeocin resistance gene functional in E.coli and the A.fumigatuspyrG gene which can be used to select transformants of filamentous fungi that lack OMP-decarboxylase (e.g. A.nidulans pyrG, Aspergillus niger pyrA, Neurospora crassa pyr4 mutants). Because pCDA21 carries an R6Kγ origin of replication, it cannot replicate in E.coli strains that do not express the pir gene (14). Therefore, amplification products obtained from this plasmid can be introduced into E.coli pir– strains without the need for DpnI treatment or gel purification that has been used to eliminate the matrix DNA (4,5). Allelic replacement on cosmids To validate the strategy outlined in Figure 1, we focused on the A.nidulans treA gene which encodes an acid trehalase required for growth of A.nidulans on trehalose: inactivation of treA can be easily scored by comparing growth of transformants on minimal medium containing glucose or trehalose as the sole carbon source (8). Cosmid W14C08 which contains the treA gene and flanking genomic regionwas introduced into an E.coli strain carrying pKOBEG. The resulting strain was transformed with (i) a 7 kb NotI fragment derived from plasmid pTRE18 and composed of a spc/pyrG marker flanked by 2.3 and 0.7 kb of DNA homologous to the 5′ and 3′ region of the treA gene, respectively or (ii) a PCR product obtained through amplification of the zeo/pyrG cassette of pCDA21 using oligonucleotides with 50 bp of homology to the 5′- or 3′-end of the treA coding region (regions F and B in Fig. 1) and 20 bp of homology in the zeo/pyrG cassette (regions PZ and PP in Fig. 1). Tens to hundreds of SpcR or ZeoR transformants were obtained in both cases. Characterization of a subset of these transformants by restriction enzyme analysis and PCR analysis showed that most resulted from the expected allelic exchange on cosmid W14C08. cTRE19 and cTRE20 (Fig. 3) are selected derivatives of W14C08 obtained using the NotI fragment of pTRE18 or PCR-amplified pCDA21, respectively. Sequencing of the region at the boundary between genomic DNA and the selective marker used for allelic replacement demonstrated exact gene replacement in cTRE20, confirming that 50 nt of homology are sufficient to promote recombination between the PCR product and a target gene located on an episome (5). Allelic replacement in A.nidulans Allelic replacement in A.nidulans is obtained using linear replacement cassettes with DNA homologous to the target locus flanking a selectable transformation marker. However, we reasoned that allelic replacement at the treA locus might occur upon transformation with circular cTRE19 or cTRE20 because of the large region of homology to the flanking regions of the treA locus that are located on both sides of the pyrG gene in these cosmids. Hence, protoplasts of A.nidulans strain FGSC773 were transformed using circular cTRE19 and cTRE20 as well as pTRE12 and pTRE18, which carry standard replacement cassettes for the treA gene cloned into pBLUESCRIPT SN+ (8,19). Approximately 25 prototrophic transformants obtained with each plasmid were tested for their ability to grow on minimal trehalose medium. The results in Table 1 show that while no treA– transformants could be obtained upon transformation of FGSC773 by pTRE12 or pTRE18, up to 29% of the transformants obtained using cTRE20 had a treA– phenotype although the frequency of allelic replacement using cTRE20 was variable. Allelic replacement in five treA– transformants obtained with cTRE20 was confirmed by PCR (data not shown) and Southern analysis (Fig. 3), thus suggesting that in contrast to plasmids carrying standard allelic replacement cassettes, circular cosmids can efficiently recombine with the A.nidulans genome to induce allelic replacement. Furthermore, Southern analysis revealed that gene replacement events were not accompanied by the integration of ectopic copies of the transforming DNA (Fig. 3). In order to increase the frequency of allelic replacement, transformation of A.nidulans FGSC773 with linearized cTRE20 was tested. First, cTRE20 was treated with two restriction enzymes that do not cleave the zeo/pyrG marker and were known to generate large DNA fragments encompassing the mutated treA allele: ClaI has several cleavage sites in W14C08, including two located at both ends of a 10 kb fragment encompassing the treA gene; SfiI cleaves W14C08 (and hence cTRE20) at only one site in the vector backbone. The results in Table 1 show that using ClaI-treated cTRE20, allelic replacements at a frequency >50% could be obtained. A higher replacement frequency (66%) was observed with SfiI-treated cTRE20 but might be considered not significant because of the small number of transformants obtained in this experiment. These replacement frequencies were significantly higher than those observed when using NotI-digested pTRE12 (5.3%; 8) or with circular cTRE20 (see above). Second, cTRE20 was treated with DNase I to an extent where only linearization of the cosmid would occur (data not shown). The results in Table 1 show that the frequency of allelic replacement increased steadily with the extent of DNase I treatment. DISCUSSION In this report, we have presented a rapid, simple and efficient methodology to generate allelic replacements in the filamentous fungus A.nidulans. This methodology takes advantage of the Redα and Redβ recombination functions of phage λ to promote in E.coli the in vivo exchange of a wild-type allele located on a cosmid by a PCR-generated mutant allele. The resulting cosmid with the mutant allele is then used to transform A.nidulans and obtain allelic exchange with the chromosomal wild-type locus (Fig. 1). The method is rapid because it does not involve a single subcloning step and only requires limited sequence data on the target gene to be able to generate mutant cosmid. Indeed, we have confirmed the previously published results of Muyrers et al. (5) who showed that 50 bp of homology between a episomally-located gene and a PCR product is sufficient to sustain recombination when Redα and Redβ (or E.coli RecE and RecT) are produced in combination with Redγ which inhibits the RecBCD exonuclease V. The method is simple because the vectors that carry the recombination functions and the zeo/pyrG bi-functional marker have been optimized to be compatible with ColE1-derived cosmids and to avoid purification and DpnI treatment of the PCR product that is used to generate the mutant cosmid. Finally, the method is efficient because allelic replacements on cosmids are obtained at high frequencies and, most interestingly, allelic replacements in A.nidulans are obtained at frequencies that are unprecedented in this fungus. In particular, we have shown that allelic replacement at the A.nidulans treA locuscould be obtained at frequencies up to 30% when using a circular recombinant cosmid while allelic exchange is never observed when a vector carrying a standard replacement cassette with 1–2 kb of homology to the target locus is used in a circular form. Furthermore, by linearizing the recombinant cosmid we have been able to increase the replacement frequency to above 50% which is significantly higher than the replacement efficiency we had observed at this locus with a standard replacement cassette (5%; 10) which was close to the 5–10% replacement efficiency commonly observed in A.nidulans and other filamentous fungi when using this type of methodology. Although linearization is not always possible because of the lack of knowledge of the genomic region flanking the target gene, the design of new cosmid libraries with a very rare cleavage site (e.g. for a meganuclease) and the increasing knowledge of genome sequence should facilitate this step. Nevertheless, we have experienced that in most instances the use of a circular cosmid is sufficient to identify the appropriate allelic replacement within a limited population of transformants (M.-K.Chaveroche and C.d’Enfent, unpublished data). Although pCDA21 has been developed for the generation of allelic replacements in A.nidulans, it can be used to create similar events in filamentous fungi where OMP-decarboxylase mutants are available (e.g. A.niger pyrA and N.crassa pyr-4 mutants). In this plasmid, we have used the A.fumigatus gene to prevent recombination in A.nidulans between the selectable marker and the endogenous pyrG89 allele. A similar cassette has been designed that uses the A.nidulans pyrG gene and can therefore be used to generate allelic replacements in A.fumigatus (M.-K.Chaveroche and C.d’Enfent, unpublished data). Furthermore, new cassettes are being developed using dominant selectable markers, in particularhygromycin resistance, which has proven to be functional in a wide variety of filamentous fungi without the need for an auxotrophic mutant (20). In this study, we have applied this two-step replacement methodology to exchange the entire open reading frame of the A.nidulans treA gene with a selectable marker. Because the site of recombination of the selectable marker at the target locus in the cosmid is precisely defined by the sequence of the oligonucleotides that are used for its amplification, other types of allelic exchanges and hence mutations can be generated using this methodology. In particular it should be possible to generate conditional mutants by promoter replacement or gene fusions provided that allelic replacement cassettes with a regulatable promoter or a reporter gene in addition to the selectable marker are available. ACKNOWLEDGEMENTS We thank K. Murphy, J. Beckwith, G. Church, H. Krish and D. Mazel for providing the different plasmids that were used in the course of this study. * To whom correspondence should be addressed. Tel: +33 1 40 61 32 57; Fax: +33 1 45 68 87 90; Email: [email protected] View largeDownload slide Figure 1. A two-step methodology for allelic exchange in A.nidulans. F and B refer to the homology extensions corresponding to regions flanking the target gene (open arrow). PZ and PP refer to priming sites on the bi-functional zeo/pyrG transformation marker. View largeDownload slide Figure 1. A two-step methodology for allelic exchange in A.nidulans. F and B refer to the homology extensions corresponding to regions flanking the target gene (open arrow). PZ and PP refer to priming sites on the bi-functional zeo/pyrG transformation marker. View largeDownload slide Figure 2. Maps of recombination plasmid pKOBEG (A) and plasmid pCDA21 (B) carrying the zeo/pyrG bi-functional transformation marker. View largeDownload slide Figure 2. Maps of recombination plasmid pKOBEG (A) and plasmid pCDA21 (B) carrying the zeo/pyrG bi-functional transformation marker. View largeDownload slide Figure 3. Allelic exchange on cosmid W14C08. (A and B) Replacement using a 7.0 kb NotI fragment derived from pTRE18, yielding cosmid cTRE19 (A) or PCR-amplified pCDA21, yielding cosmid cTRE20 (B). Solid line, A.nidulans DNA; open box, spectinomycin-resistance gene (A) or zeocin-resistance gene (B); gray box, A.fumigatus pyrG gene; E, EcoRI; P, PmlI; N, NotI; arrow, A.nidulans treA coding region; open circle, treA start codon; solid circle, treA stop codon; double-arrow, deletion in the treA coding region resulting from the allelic exchange; arrowheads, 50 bp homology extensions present in the composite primers used to amplify the zeo/pyrG bi-functional transformation marker. The size of the EcoRI restriction fragments in the cosmids, pTRE18/NotI fragment and PCR product are indicated in kb. (C) Ethidium bromide-stained gel of EcoRI-digested W14C08, cTRE19 and cTRE20. The 8.0 kb fragment of W14C08 and the resulting 5.0 and 4.7 kb fragments in cTRE19 and cTRE20, respectively, are indicated by solid arrowheads. The 2.1 and 0.5 kb fragments corresponding to the spc (cTRE19) and zeo (cTRE20) genes, respectively, are indicated by open arrowheads. The 1.9 kb fragment corresponding to the A.fumigatus pyrG geneis indicated by a grey arrowhead. MWM, molecular weight markers, 1 kb ladder (Gibco BRL). Representative molecular weights are indicated in kb. (D) Southern hybridization of EcoRI-digested genomic DNA of A.nidulans FGSC773 (wild-type, lane 1) and of seven transformants: two treA+ transformants with ectopic integrations of pTRE20 (lanes 2 and 3) and five treA– transformants with an appropriate gene replacement at the treA locus (lanes 4–8). Genomic DNA was transferred onto nylon membranes and probed for treA (top) or a control single-copy gene (bottom). An 8.0 kb fragment is detected when treA is intact while a 4.7 kb fragment is detected when the treAΔ::(zeo/pyrG) allele is present. Similar intensities of the hybridization signals obtained using the treA and control probes demonstrates that the treAΔ::(zeo/pyrG) allele is present in a single copy. View largeDownload slide Figure 3. Allelic exchange on cosmid W14C08. (A and B) Replacement using a 7.0 kb NotI fragment derived from pTRE18, yielding cosmid cTRE19 (A) or PCR-amplified pCDA21, yielding cosmid cTRE20 (B). Solid line, A.nidulans DNA; open box, spectinomycin-resistance gene (A) or zeocin-resistance gene (B); gray box, A.fumigatus pyrG gene; E, EcoRI; P, PmlI; N, NotI; arrow, A.nidulans treA coding region; open circle, treA start codon; solid circle, treA stop codon; double-arrow, deletion in the treA coding region resulting from the allelic exchange; arrowheads, 50 bp homology extensions present in the composite primers used to amplify the zeo/pyrG bi-functional transformation marker. The size of the EcoRI restriction fragments in the cosmids, pTRE18/NotI fragment and PCR product are indicated in kb. (C) Ethidium bromide-stained gel of EcoRI-digested W14C08, cTRE19 and cTRE20. The 8.0 kb fragment of W14C08 and the resulting 5.0 and 4.7 kb fragments in cTRE19 and cTRE20, respectively, are indicated by solid arrowheads. The 2.1 and 0.5 kb fragments corresponding to the spc (cTRE19) and zeo (cTRE20) genes, respectively, are indicated by open arrowheads. The 1.9 kb fragment corresponding to the A.fumigatus pyrG geneis indicated by a grey arrowhead. MWM, molecular weight markers, 1 kb ladder (Gibco BRL). Representative molecular weights are indicated in kb. (D) Southern hybridization of EcoRI-digested genomic DNA of A.nidulans FGSC773 (wild-type, lane 1) and of seven transformants: two treA+ transformants with ectopic integrations of pTRE20 (lanes 2 and 3) and five treA– transformants with an appropriate gene replacement at the treA locus (lanes 4–8). Genomic DNA was transferred onto nylon membranes and probed for treA (top) or a control single-copy gene (bottom). An 8.0 kb fragment is detected when treA is intact while a 4.7 kb fragment is detected when the treAΔ::(zeo/pyrG) allele is present. Similar intensities of the hybridization signals obtained using the treA and control probes demonstrates that the treAΔ::(zeo/pyrG) allele is present in a single copy. Table 1. Allelic replacement at the treA locus using circular or linearized recombinant cosmids Plasmid/cosmid  Experimenta  Homology to treA locusb  treA–/totalc  Circular        pTRE12  1  2.3/1.2  0/25  pTRE18  1  2.3/1.2  0/25  cTRE19  1  ND  2/24  cTRE20  1  ND  5/24  cTRE20  2  ND  8/24  cTRE20  3  ND  0/24  Linearized         pTRE12/NotI  Ref. 10  2.3/0.7  4/75  cTRE20/ClaI  2  ND  13/25  cTRE20/SfiI  2  ND  2/3  cTRE20/DNase I (1 min)  3  ND  1/25  cTRE20/DNase I (2 min)  3  ND  3/25  cTRE20/DNase I (5 min)  3  ND  6/25  cTRE20/DNase I (10 min)  3  ND  2/20  cTRE20/DNase I (15 min)  3  ND  8/24  Plasmid/cosmid  Experimenta  Homology to treA locusb  treA–/totalc  Circular        pTRE12  1  2.3/1.2  0/25  pTRE18  1  2.3/1.2  0/25  cTRE19  1  ND  2/24  cTRE20  1  ND  5/24  cTRE20  2  ND  8/24  cTRE20  3  ND  0/24  Linearized         pTRE12/NotI  Ref. 10  2.3/0.7  4/75  cTRE20/ClaI  2  ND  13/25  cTRE20/SfiI  2  ND  2/3  cTRE20/DNase I (1 min)  3  ND  1/25  cTRE20/DNase I (2 min)  3  ND  3/25  cTRE20/DNase I (5 min)  3  ND  6/25  cTRE20/DNase I (10 min)  3  ND  2/20  cTRE20/DNase I (15 min)  3  ND  8/24  aThree independent experiments were performed. bLength in kb of the DNA homologous with the treA locus located on each side of the selectable marker in pTRE12 and pTRE18. These values are not known but >5 kb for cTRE19 and cTRE20. ND, not determined. cNumber of transformants with a treA– phenotype relative to the total number of transformants tested. The treA phenotype was scored on minimal trehalose medium. View Large References 1 van den Hondel,C.A.M.J.J. and Punt,P.J. ( 1991) In Peberdy,J.F., Caten,C.E., Ogden,J.E. and Bennett,J.W. (eds), Applied Molecular Genetics of Fungi. Cambridge University Press, Cambridge, UK, pp. 1–28. Google Scholar 2 d’Enfert,C., Weidner,G. and Brakhage,A.A. ( 1999) In Brakhage,A.A., Jahn,B. and Schmidt,A. (eds), Aspergillus fumigatus: Biology, Clinics and Molecular Approaches to Pathogenicity. S. Karger AG, Basel, Switzerland, Vol. 2, pp. 149–166. Google Scholar 3 Murphy,K.C. ( 1998) J. Bacteriol. , 180, 2063–2071. Google Scholar 4 Zhang,Y., Buchholz,F., Muyrers,J.P.P. and Stewart,A.F. ( 1998) Nature Genet. , 20, 123–128. Google Scholar 5 Muyrers,J.P.P., Zhang,Y., Testa,G. and Stewart,A.F. ( 1999) Nucleic Acids Res. , 27, 1555–1557. Google Scholar 6 Datsenko,K.A. and Wanner,B.L. ( 2000) Proc. Natl Acad. Sci. USA , 97, 6640–6645. Google Scholar 7 Guzman,L.M., Belin,D., Carson,M.J. and Beckwith,J. ( 1995) J. Bacteriol ., 177, 4121–4130. Google Scholar 8 d’Enfert,C. and Fontaine,T. ( 1997) Mol. Microbiol. , 24, 203–216. Google Scholar 9 Sambrook,J., Fritsch,E.F. and Maniatis,T. ( 1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Google Scholar 10 Ausubel,F.M., Brent,R., Kingston,R.E., Moore,D.D., Seidman,J.G., Smith,J.A. and Struhl,K. (eds) ( 1992) Short Protocols in Molecular Biology. John Wiley & Sons, New York, NY. Google Scholar 11 Hanahan,D., Jessee,J. and Bloom,F.R. ( 1991) Methods Enzymol. , 204, 63–113. Google Scholar 12 Link,A.J., Phillips,D. and Church,G.M. ( 1997) J. Bacteriol. , 179, 6228–6237. Google Scholar 13 Prentki,P. and Krisch,H.M. ( 1984) Gene , 29, 303–313. Google Scholar 14 Miller,V.L. and Mekalanos,J.J. ( 1988) J. Bacteriol ., 170, 2575–2583. Google Scholar 15 Weidner,G., d’Enfert,C., Koch,A., Mol,P.C. and Brakhage,A.A. ( 1998) Curr. Genet. , 33, 378–385. Google Scholar 16 Osmani,S.A., May,G.S. and Morris,R.N. ( 1987) J. Cell Biol. , 104, 1495–1504. Google Scholar 17 Prade,R.A., Griffith,J., Kochut,K., Arnold,J. and Timberlake,W.E. ( 1997) Proc. Natl Acad. Sci. USA , 94, 14564–14569. Google Scholar 18 Brody,H., Griffith,J., Cuticchia,A.J., Arnold,J. and Timberlake,W.E. ( 1991) Nucleic Acids Res. , 19, 3105–3109. Google Scholar 19 d’Enfert,C., Diaquin,M., Delit,A., Wuscher,N., Debeaupuis,J.-P., Huerre,M. and Latgé,J.-P. ( 1996) Infect. Immunol. , 64, 4401–4405. Google Scholar 20 Punt,P.J. and van den Hondel,C.A. ( 1992) Methods Enzymol. , 216, 447–457. Google Scholar

Penchovsky, Robert;Birch-Hirschfeld, E.;McCaskill, John S.

doi: 10.1093/nar/28.22.e98pmid: 11071952

Abstract A novel approach for light-dependent covalent immobilisation of synthetic DNA oligomers to amino-coated paramagnetic beads is described. A hetero-bifunctional photo-reactive cross-linking chemical, 4-nitrophenyl 3-diazopyruvate, is applied to attach 5′ amino-modified DNA to both silica and polystyrene paramagnetic beads. The coupling yields are comparable with similar methods in which no photo-reactive chemicals are used. The immobilised DNA on the polystyrene and silica beads was used efficiently in hybridisation experiments. An extension of this approach to light-directed immobilisation of specific DNA to beads, located at different positions in micro-flow reactors, opens up a range of integrated applications to complex diagnostics, evolutionary biotechnology and novel areas such as DNA computing. Received July 14, 2000; Revised and Accepted September 25, 2000. INTRODUCTION In recent years, DNA chip-based assays have become a familiar approach suitable for a broad range of applications such as expression analysis (1,2), polymorphism analysis and genotyping (3,4), and the detection of pathogens (5). The expansion of DNA chip technology has encouraged the rapid development of light-directed oligonucleotide synthesis (6,7). Light-directed oligonucleotide synthesis has proven to be an efficient method for fabricating probe arrays with densities as high as 106 unique sequences/cm2 (8). Nevertheless, there are some common errors in optical oligonucleotide synthesis, such as premature truncations of the growing strand and base deletions, which seem difficult to avoid. Potentially, the error rate in DNA chips due to these factors could be reduced by replacing the light-directed oligonucleotide synthesis with light-directed oligonucleotide immobilisation. Despite the existence of a great number of different chemical methods for specific 5′- and 3′-end covalent DNA attachment, there are only a few approaches for the photo-immobilisation of DNA (9). In this article, we propose a novel method for light-dependent covalent immobilisation of 5′-end amino-modified single-stranded DNA, based on the hetero-bifunctional, photo-reactive cross-linking agent 4-nitrophenyl 3-diazopyruvate (DAPpNP). As a solid support for photo-immobilisation of DNA, we employ silica and polystyrene amino-coated super-paramagnetic beads. Our goal is to apply this photo-dependent DNA immobilisation to allow a programmed immobilisation of DNA to beads at specific sites in micro-flow reactors. This patterned immobilisation can then direct the sequence specific selection from complex DNA populations (J.S.McCaskill, submitted) and thereby enable an optical specification of different problem instances in the field of DNA computing (10). A super-paramagnetic bead-based module for integrated complex selection in micro-flow reactors has been described (J.S.McCaskill, submitted) together with a sample application (to the maximal clique problem) designed to make use of the current light-directed immobilisation procedure. In contrast to chemical cross-linking reagents, which have been successfully applied for DNA immobilisation (11,12), photo-reactive cross-linking reagents have primarily been employed as tools for defining interactions among proteins, nucleic acids, ligands and their receptors (13,14). The major disadvantage in using photo-reactive cross-linking reagents for DNA immobilisation is that such agents form extremely reactive groups after photo-activation, usually causing many non-specific side reactions (9). In this work, for the attachment of DNA we avoid directly utilising the amino-ketene group, formed after photolysis of a diazo group of diazopyruvic acid, because of its high reactivity and its instability in aqueous solutions. Instead, we facilitate the transformation of the ketene group into a carboxyl group, which is then utilised for reaction with a 5′ amino-modified deoxynucleotide oligomer in the presence of carbodiimide. MATERIALS AND METHODS Paramagnetic beads and oligodeoxynucleotides Silica and polystyrene super-paramagnetic beads were purchased from Micromod GmbH (Rostock, Germany). All beads are essentially monodisperse. The properties of the beads are presented in Table 1. Oligomers were obtained from IBA-NAPS (Göttingen, Germany). The DNA was 5′ amino-labelled using a C6 linker. All DNA was purified to HPLC grade. The oligomer sequences used are shown in Table 2. Modification of amino-coated beads with the photo-reactive bifunctional cross-linker DAPpNP DAPpNP was obtained from Molecular Probes (Eugene, OR). The cross-linker was dissolved to produce a 200 mM solution in dry DMSO. 100 µl of this solution was added to 5 mg amino-coated beads resuspended in 400 µl dry DMSO. The reaction was performed for 2 h at room temperature with continuous shaking in the dark (see Fig. 1, reaction 1). The beads were washed several times in the dark to remove any traces of nitrophenol with deionized H2O until there was no longer any measurable absorbance at 412 nm in the rinsing solution. Photolysis of the diazo group to amino ketene and its transformation to the carboxyl group The diazo group attached to beads (as above) was photolysed by exposure to UV radiation for 30 s applying a mercury lamp with a power of 15 W from 280 to 315 nm and 75 W from 315 to 400 nm (UV-F400, Panacol-Elosol GmbH, Oberursel, Germany). In the case of immobilisation of amino-modified DNA applying 1-ethyl-3-(3-dimethyl-aminopropyl) carbodiimide (EDAC) (Sigma) (see Fig. 1, left-hand route) 5 mg beads were placed into a quartz cuvette in 500 µl deionized H2O. The reaction mix was incubated for 0.5 h at room temperature while continuously shaking in the dark (see Fig. 1, reactions 2 and 4). In the case of direct immobilisation of amino-modified oligomers (see Fig. 1, right-hand route), 25 µl (100 pmol/µl) of the 5′ amino-modified 35 nt oligomer was added. The reaction was incubated for 2 h at room temperature with continuous shaking in the dark. Negative controls were performed, as for the reactions described above, but in the dark without exposure to radiation. Covalent attachment of 5′ amino-modified DNA oligomers to a photo-induced carboxyl group An aliquot (2.5 nmol) of the 35 nt 5′ amino-modified oligomer was used for the covalent linkage with a photo-induced carboxyl group on 5 mg beads dissolved in 500 µl deionized H2O in the presence of 10 mM EDAC. The reaction was performed for 3 h at room temperature with continuous shaking in the dark (see Fig. 1, reaction 5). In order to remove non-covalently bound DNA, the beads were treated with SPSC buffer (50 mM sodium phosphate, 1 M NaCl, pH 6.5) for 1.5 h at 42°C, and then with 100 mM Tris–acetate pH 8.0 for 1 h at 42°C in the dark. Quantification of the DNA attached to the beads The peptide bond formed between the immobilised DNA and the bead spacer was destroyed with 28% NH4OH (Sigma). The reaction was performed at 48°C for 3 h with continuous shaking. NH4OH was removed by evaporation with a SpeedVac Concentrator (Savant Instrument, Inc., Holbrook, NY). The cleaved DNA was purified on a NAP-10 column (Pharmacia Biotech, Uppsala, Sweden) as described by the manufacturer. The concentration of DNA cleaved from the beads was estimated by UV spectroscopy with a Cary 3E photometer (Varian Inc., Walnut Creek, USA) at 260 nm as described by Sambrook et al. (15). HPLC analysis HPLC analysis was performed on an HPLC liquid chromatograph (Shimadzu LC-10AT with system controller SCL-10A and UV-vis detector SPD-10 Avi at 260 nm) using the software package Class VP v.4.0 (Shimadzu, Japan). An anion-exchange column (mono-Q HR5/5, Pharmacia) was loaded with a linear salt gradient (bufferA: 10 mM NaOH in H2O; buffer B: 1 M NaCl in 10 mM NaOH) at a flow rate of 1.5 ml/min, increasing buffer B from 0 to 100% in 20 min. DNA hybridisation on beads and its detection Beads (2.5 mg) with immobilised DNA (see Table 2, row 3) were resuspended in 500 µl 100 mM Tris–acetate buffer pH 8.0 in the presence of 2 µM 5′ rhodamine 6G labelled DNA (see Table 2, row 4). The hybridisation reactions took place at 42°C for 1 h while continuously shaking in the dark, after which the hybridisation solution was removed and the beads were washed three times with 500 µl 100 mM Tris–acetate buffer pH 8.0 at 42°C for 15 min. Finally, the beads were resuspended in 100 µl water and incubated at 95°C for 2 min. The fluorescent signal of the denatured DNA in the supernatant was measured by a spectrofluorimeter with excitation at 524 nm and emission measured at 557 nm (FluorMax-2, Instruments S.A., Inc., Edison, NJ). Light-directed DNA immobilisation to beads in a micro-flow reactor In order to demonstrate a spatially defined light-directed DNA immobilisation the beads were incorporated in a micro-flow reactor. The design and the fabrication of the micro-reactor are described in McCaskill et al. (16). The micro-reactor was anodically bonded to two 500-µ-thick pyrex (borosilicate) wafers, where 1 mm of the wafer has a 50% transmission at 306 nm. The reactor has two inlet and two outlet channels (see Fig. 3C). Both inlet channels are filled with polystyrene amino-coated beads (see Table 1, row 2) in the dark. The beads are restrained in inlet channels by a ledged bead barrier. They were previously treated with a solution of DAPpNP and washed as described above. Water was pumped with a flow rate of 2 µl/min via both inlet channels of the micro-flow reactor in the dark using a precision syringe pump model 260 manufactured by World Precision Instruments Inc. (Sarasota, FL). The reactor was placed under a UV lamp with part of one of the inlet channels covered by an opaque foil (see Fig. 3B). The micro-flow reactor was irradiated by the lamp twice for 30 s. The aqueous solution was replaced after 2 h by a solution of 10 mM EDAC and 2.5 nmol of the 35 nt 5′ amino-modified and 3′ fluorescein-modified oligomer (see Table 2, row 3). The reaction was performed for 2 h at a pump rate of 1 µl/min. In order to remove as much of the non-covalently attached DNA as possible, the micro-flow reactor was washed three times with 1 ml 50% DMSO and three times with 1 ml 100 mM Tris–acetate pH 8.0 buffer at a 2 µl/min flow rate. Spatially resolved detection of DNA immobilisation to beads A CCD camera model LN/1024 TKB/1 (Princeton Instrument Inc., Monmouth Junction, NJ) was employed to capture images from the micro-flow reactors. The illumination was performed either by white light or by laser beam at 514.5 nm from an argon-ion laser model 2080-15S manufactured from Spectra-Physics Lasers, Inc. (Mountain View, CA). A homogenous pool of light of diameter ~10 cm was created by acoustic vibration of a multi-mode optical fibre at 80 Hz. An interference filter was placed between the camera and the sample, custom designed from AHF-Analysentechnik (Tübingen, Germany) with an OD of 6 at 514 nm and transmission of 85–90% at 523–600 nm. For data acquisition and image processing, a customised version of the software PMIS v.2.0.1 (GKR Computer Consulting, Boulder, CO) was employed. RESULTS AND DISCUSSION The overall scheme of the photo-dependent immobilisation is shown in the left-hand route of Figure 1. The 4-nitrophenyl ester group of DAPpNP reacts with amines producing diazopyruvic acid amides that undergo UV photolysis to generate ketene amides through carbene formation and Wolf rearrangement (17,18). The ketene group is unstable in water, reacting readily to form a carboxyl group (17). This photo-induced carboxyl group can be used for a covalent immobilisation of 5′- or 3′-end amino-labelled DNA in the presence of EDAC, via the formation of a peptide bond (11). An alternative, slightly more involved, mechanism shown in the right-hand route of Figure 1 has also been investigated. It starts with the same photo-activated ketene group, but coupling occurs directly with 5′-end amino-modified DNA. Chemical stability of the diazo group The key point in our photo-immobilisation strategy is the chemical stability of the diazo group. Diazomethane and other diazoalkyl derivatives have been used in the HPLC analysis of low molecular weight compounds to label carboxyl groups of fatty acids (19). Diazoalkanes and diazoacetyl compounds (amides and esters) react with carboxylate groups even without catalysts. This should not be a problem here because both carboxyl and diazo groups are attached to the bead surface spacers (see Table 1) so that their reaction should be limited by geometric constraints. The diazo compound was found to be stable in water at room temperature (20). Because this point was critical for the current method, we tested the stability of the diazo compound in the presence of EDAC and 5′ amino-modified DNA oligomer in water, using an HPLC analysis as described in Materials and Methods. First, we examined the retention time of a 5′ amino-modified oligomer (see Table 1, oligomer 1) in deionized H2O using the HPLC system. A retention time of 11.9 min for this DNA oligomer was found (peak UV absorption). Secondly, 40 mM of DAPpNP with a 2 µM aqueous solution of the same DNA in the presence of 100 mM EDAC were incubated at room temperature for 2 h and for 8 h in the dark. An identical HPLC analysis showed that the retention time of the DNA was unchanged for both incubation times, 11.9 min (data not shown). To ascertain the sensitivity of this measurement to potential reactions of the diazo group, we repeated the experiment in a buffer solution in which it proved reactive. We incubated 40 mM of DAPpNP with a 2 µM solution of the DNA oligomer in 100 mM borate buffer pH 8.6 for 2 and 8 h. For both incubation times, two peaks of DNA were observed, at 11.9 and 12.5 min (data not shown). The second peak indicates a reaction between the 5′ amino group of DNA and DAPpNP, resulting in the production of diazopyruvic acid amides at the 5′-end of the DNA oligomer. Since such reactive products can be discriminated by HPLC analysis, we can conclude from the experiment above that the diazo-compound is stable in water in the presence of EDAC and amino-modified DNA at room temperature for at least 8 h in the dark. Photo-dependent immobilisation of DNA oligomers to amino-coated paramagnetic beads The immobilisation reaction was performed as described in Materials and Methods together with negative controls. Amino-coated silica and polystyrene beads, 15 µm in diameter, were used as a solid support in the first set of experiments. The DNA products were cleaved from the beads and were purified as described in Materials and Methods. The DNA products were quantified by UV absorbance as reported in Table 3. The negative control reaction 2 (see Fig. 2 and Table 3, column 2) was performed in the same way as the positive reaction 1 (see Fig. 2 and Table 3, column 1), but without exposure to UV radiation. Negative control 3 (see Fig. 2 and Table 3, column 3) was made without treatment of beads with DAPpNP negative control 4 (see Fig. 2 and Table 3, column 4) was made with neither DAPpNP nor EDAC and negative control 5 (see Table 3, column 5) was made without EDAC. As one can see from Figure 2 and Table 3, the non-specific immobilisation in all negative controls is the same, so that the non-specific immobilisation is due to surface interaction but not to chemical reactions. We demonstrated a spatially defined light-directed DNA immobilisation to magnetic beads incorporated in a micro-flow reactor. As one can see from the fluorescent image in Figure 3B, the beads in the channel not exposed to radiation show a very low fluorescent signal in comparison with the fluorescent signal from the beads exposed to radiation. Both inlet channels of the micro-flow reactor are indeed filled with beads as shown in the white light image of Figure 3A. The quantification of the fluorescent signal is shown in Figure 4. Comparison of various beads and surface coatings for immobilisation Carboxyl-coated beads may be used directly for immobilisation of amino-modified DNA using EDAC (without the above photo-activation technique). A comparison with such direct immobilisation provides a measure of the efficiency of the photo-immobilisation procedure introduced above. To implement the above efficiency comparison, the yield of our photo-immobilisation strategy on polystyrene amino-coated beads as described in the previous section was compared with the DNA immobilisation yield on polystyrene carboxyl-coated beads of the same size (15 µm) but applying only a one-step reaction with 5′ amino-coated 35 nt DNA oligomer in the presence of EDAC. As can be seen from Table 4, the immobilisation yield of the photo-immobilisation procedure (210 pmol/mg beads) is even higher than the immobilisation yield from the well-known EDAC synthesis (150 pmol/mg beads). This is due to less non-specific immobilisation on carboxyl-coated beads than on amino-coated ones. Hybridisation analysis Hybridisation analyses were made in order to estimate the efficiency of the applied immobilisation methods on carboxyl- and amino-coated polystyrene and silica beads. A 35 nt DNA oligomer (see Table 2, oligomer 3) was immobilised on carboxyl-coated polystyrene beads as described in Materials and Methods. Immobilised DNA on 2.5 mg beads was hybridised with the complementary 5′ rhodamine-labelled 27 nt oligomer (see Table 2, oligomer 4) and measured by fluorescence as described in Materials and Methods. As one can see in Figure 2, the hybridisation yield on amino-coated silica beads after applying the photo-dependent immobilisation is ~25% less in comparison with the hybridisation yield on carboxyl-coated silica beads after direct immobilisation of 5′ amino-modified DNA by EDAC. At the same time, the background signal is almost the same (with the exception of amino-coated polystyrene beads after applying the photo-dependent immobilisation) despite the fact that silica beads showed more non-specific DNA attachment. This could imply that the non-specifically attached DNA is not accessible to hybridisation. CONCLUSIONS We have shown that the hetero-bifunctional photo-reactive cross-linking agent DAPpNP can be employed for light-directed immobilisation of DNA via a photo-induced formation of a carboxyl group and final immobilisation of 5′- or 3′-end amino-modified synthetic DNA in the presence of EDAC. This photo-dependent DNA immobilisation strategy for light-directed attachment of DNA oligomers to paramagnetic beads may be applied to parallel investigations of DNA hybridisation in micro-flow reactors (J.S.McCaskill, submitted). Beads placed in different positions in micro-flow reactors may be selectively addressed (i.e. loaded with oligomers by photo-dependent DNA immobilisation) using a photo-lithographic mask and analysed individually in parallel. We expect the technique to further extend our ability to perform parallel in vitro analyses with DNA. ACKNOWLEDGEMENTS The authors wish to thank Andreas Opitz for helpful discussions, Frank-Ulrich Gast for critically reading the manuscript, Thomas Kirner and Harald Mathis for a set-up of the laser detection system and Marlies Gohlke for producing the micro-flow reactors. The first few months of this work were supported at the Institute for Molecular Biotechnology in Jena, Germany. * To whom correspondence should be addressed. Tel: +49 2241 141 517; Fax: +49 2241 141 511; Email: [email protected] View largeDownload slide Figure 1. Overall scheme of photo-immobilisation. Left-hand route: the main photo-immobilisation scheme using the zero-crosslinker EDAC to attach 5′ amino-modified DNA oligomers to beads. (1) The 4-nitrophenyl ester group of DAPpNP reacts with amines on the beads producing diazopyruvic. (2) Photolysis of the diazo group attached to the beads by UV illumination and (3) formation of ketene amides with (4) conversion of the ketene group to carboxyl in water. (5) Covalent immobilisation of 5′ amino-modified oligomer to the photo-induced carboxyl group (attached to the beads) in the presence of EDAC via peptide-bond formation. Right-hand route: (6) Direct utilisation of the amino-ketene group for immobilisation of 5′ amino-modified DNA oligomers. View largeDownload slide Figure 1. Overall scheme of photo-immobilisation. Left-hand route: the main photo-immobilisation scheme using the zero-crosslinker EDAC to attach 5′ amino-modified DNA oligomers to beads. (1) The 4-nitrophenyl ester group of DAPpNP reacts with amines on the beads producing diazopyruvic. (2) Photolysis of the diazo group attached to the beads by UV illumination and (3) formation of ketene amides with (4) conversion of the ketene group to carboxyl in water. (5) Covalent immobilisation of 5′ amino-modified oligomer to the photo-induced carboxyl group (attached to the beads) in the presence of EDAC via peptide-bond formation. Right-hand route: (6) Direct utilisation of the amino-ketene group for immobilisation of 5′ amino-modified DNA oligomers. View largeDownload slide Figure 2. Hybridisation analysis of DNA to bead-bound oligomers. Filled bars, total hybridisation yield; unfilled bars, background hybridisation. Sample 1, carboxyl-coated polystyrene beads: immobilisation using the zero-crosslinker EDAC. Sample 2, carboxyl-coated silica beads: immobilisation using the zero-crosslinker EDAC. Sample 3, amino-coated silica beads: photo-immobilisation using the zero-crosslinker EDAC. Sample 4, amino-coated polystyrene beads: photo-immobilisation scheme using the zero-crosslinker EDAC. Sample 5, amino-coated silica beads: direct utilisation of the amino-ketene group for immobilisation of 5′ amino-modified DNA oligomer (see Materials and Methods). View largeDownload slide Figure 2. Hybridisation analysis of DNA to bead-bound oligomers. Filled bars, total hybridisation yield; unfilled bars, background hybridisation. Sample 1, carboxyl-coated polystyrene beads: immobilisation using the zero-crosslinker EDAC. Sample 2, carboxyl-coated silica beads: immobilisation using the zero-crosslinker EDAC. Sample 3, amino-coated silica beads: photo-immobilisation using the zero-crosslinker EDAC. Sample 4, amino-coated polystyrene beads: photo-immobilisation scheme using the zero-crosslinker EDAC. Sample 5, amino-coated silica beads: direct utilisation of the amino-ketene group for immobilisation of 5′ amino-modified DNA oligomer (see Materials and Methods). View largeDownload slide Figure 3. Light-dependent DNA immobilisation to beads in a micro-flow reactor. (A) White light image. Both inlet channels of a micro-reactor are filled with beads. (B) Fluorescence image. A fluorescent signal from the beads incorporated in a micro-flow reactor. The greyed column indicates the area profiled in Figure 4 and shows the line of segregation between the irradiated part (on the left-hand side) and non-irradiated part (on the right-hand side) of the inlet channel on the top of the picture. (C) Micro-flow reactor scheme. The two inlet channels (on the right-hand side of the scheme), two outlet channels (on the left-hand side of the scheme) and the bead barrier are shown. The dashed square shows the part of the micro-flow reactor presented in (A) and (B). View largeDownload slide Figure 3. Light-dependent DNA immobilisation to beads in a micro-flow reactor. (A) White light image. Both inlet channels of a micro-reactor are filled with beads. (B) Fluorescence image. A fluorescent signal from the beads incorporated in a micro-flow reactor. The greyed column indicates the area profiled in Figure 4 and shows the line of segregation between the irradiated part (on the left-hand side) and non-irradiated part (on the right-hand side) of the inlet channel on the top of the picture. (C) Micro-flow reactor scheme. The two inlet channels (on the right-hand side of the scheme), two outlet channels (on the left-hand side of the scheme) and the bead barrier are shown. The dashed square shows the part of the micro-flow reactor presented in (A) and (B). View largeDownload slide Figure 4. Quantification of the fluorescent signal of spatially defined light-directed DNA immobilisation to beads in a micro-flow reactor. An intensity section of Figure 3B is shown. Between rows 75 and 125: the signal from the beads in the channel not exposed to radiation with an intensity of 80 counts per second (c.p.s.). Between row 325 and 375: the signal from beads in the channel exposed to radiation with intensity >250 c.p.s. The background signal is ~50 c.p.s. View largeDownload slide Figure 4. Quantification of the fluorescent signal of spatially defined light-directed DNA immobilisation to beads in a micro-flow reactor. An intensity section of Figure 3B is shown. Between rows 75 and 125: the signal from the beads in the channel not exposed to radiation with an intensity of 80 counts per second (c.p.s.). Between row 325 and 375: the signal from beads in the channel exposed to radiation with intensity >250 c.p.s. The background signal is ~50 c.p.s. Table 1. Properties of the paramagnetic beads used (as specified by the manufacturer, see Materials and Methods) Bead type  Size(µm)  Content of magnetite (%)  Functional groups on the surface  Particle charge density (nmol/mg)  Carboxyl polystyrene-co-maleic   15 ± 2  15–20  Acrylic acid  6.0  Amino polystyrene-co-maleic  15 ± 2  15–20  (CH2)4NH2  3.1  Amino silica   15 ± 5  20  (CH2)3NH2  3.26  Carboxyl silica  15 ± 5  20  Citric acid  2.1  Bead type  Size(µm)  Content of magnetite (%)  Functional groups on the surface  Particle charge density (nmol/mg)  Carboxyl polystyrene-co-maleic   15 ± 2  15–20  Acrylic acid  6.0  Amino polystyrene-co-maleic  15 ± 2  15–20  (CH2)4NH2  3.1  Amino silica   15 ± 5  20  (CH2)3NH2  3.26  Carboxyl silica  15 ± 5  20  Citric acid  2.1  View Large Table 2. DNA oligonucleotides employed DNA  Type of modification  Sequence 5′–3′  Length (nt)  1  5′ Amino C6  TCCCGAAAATACTAAAAAAGCA  22  2  5′ Amino C6  TTCCCGGACGGTCACAGCTTGTCTG  25  3  5′ Amino C6 or 5′ amino C6 and3′ fluorescein  TTTTTTTTCAGACAAGCTGTGTCCGTCTCCCGGGA  35  4  5′ Rhodamine 6G or 5′ amino C6  TCCCGGGAGACGGACACAGCTTGTCTG  27  DNA  Type of modification  Sequence 5′–3′  Length (nt)  1  5′ Amino C6  TCCCGAAAATACTAAAAAAGCA  22  2  5′ Amino C6  TTCCCGGACGGTCACAGCTTGTCTG  25  3  5′ Amino C6 or 5′ amino C6 and3′ fluorescein  TTTTTTTTCAGACAAGCTGTGTCCGTCTCCCGGGA  35  4  5′ Rhodamine 6G or 5′ amino C6  TCCCGGGAGACGGACACAGCTTGTCTG  27  View Large Table 3. Yields for photo-immobilisation of DNA to silica paramagnetic beads Conditions  1. UV irradiation  2. Dark  3. No DAPpNP  4. No EDAC nor DAPpNP  5. No EDAC  DNA concentration in pmol/mg beads  460 ± 75  76 ± 12  81 ± 13  77 ± 12  81 ± 13  Conditions  1. UV irradiation  2. Dark  3. No DAPpNP  4. No EDAC nor DAPpNP  5. No EDAC  DNA concentration in pmol/mg beads  460 ± 75  76 ± 12  81 ± 13  77 ± 12  81 ± 13  View Large Table 4. Comparison of suitability of different super-paramagnetic beads for specific immobilisation of DNA Bead type  DNA immobilisation (pmol DNA/mg beads)  Non-specific DNA attachment (pmol DNA/mg beads)  Non-specific DNA immobilisation (%)  Carboxyl polystyrene-co-maleic   170 ± 20  15 ± 3  9  Amino polystyrene-co-maleic  210 ± 40  50 ± 9  24  Bead type  DNA immobilisation (pmol DNA/mg beads)  Non-specific DNA attachment (pmol DNA/mg beads)  Non-specific DNA immobilisation (%)  Carboxyl polystyrene-co-maleic   170 ± 20  15 ± 3  9  Amino polystyrene-co-maleic  210 ± 40  50 ± 9  24  View Large References 1 Gray,N.S., Wodicka,L., Thunnissen,A.W., Norman,T.C., Kwon,S., Espinoza,F.H., Morgan,D.O., Barnes,G., LeClerc,S., Meijer,L., Kim S., Lockhart,D. and Schultz,P. ( 1998) Science , 281, 533–537. 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Leiden Center for Natural Computing, The Netherlands, pp. 239–246. Google Scholar 17 Zeller,K., Meier,H., Kohlshorn,H. and Mueller,E. ( 1972) Chem. Ber. , 105, 1875–1886. Google Scholar 18 Goodfellow,V.S., Settineri,M. and Lawton,R.G. ( 1989) Biochemistry , 28, 6346–6360. Google Scholar 19 Dermar,J.C., Disher,R.M. and Wensel,T.G. ( 1992) FASEB J. , 6, A81–A81. Google Scholar 20 Friedman,A.L., Ismagilowa,G.S., Salesov,V.S. and Nivikov,S.S., ( 1972) Uspechi Chimii , 41, 722–757. Google Scholar

Schramm, Guido;Bruchhaus, Iris;Roeder, Thomas

doi: 10.1093/nar/28.22.e96pmid: 11071950

Abstract A novel approach for the amplification of cDNA ends is described. It requires only minimal amounts of material, a simple cDNA synthesis reaction and a single PCR reaction to amplify the desired 5′- or 3′-ends of a certain cDNA of interest. It combines the so called CapFinder approach with solid phase cDNA synthesis, thus almost eliminating background problems usually associated with 5′-RACE protocols. This approach could be used to generate complete 5′-ends of numerous cDNAs using only one cDNA synthesis reaction. In combination with LA PCR, several kilobases of unknown 5′-ends could be amplified. It is easy to perform, quick, inexpensive and reliable, which should enable it to replace most currently used 5′-RACE protocols. Received June 29, 2000; Revised and Accepted September 26, 2000. INTRODUCTION The extensive use of PCR approaches for the identification of novel genes requires the identification of those regions of the gene that are not enclosed by known sequences. These 3′- and 5′-regions could either be identified using conventional screening of cDNA libraries or using 5′- and 3′-RACE (rapid amplification of cDNA ends) approaches. 5′-RACE was made possible by tagging the 5′-end of a cDNA by means of different methods (1–3). Most approaches, such as homopolymeric tailing or ligation anchored tailing require a set of enzymatic reactions after completion of first strand cDNA synthesis (1). Each enzymatic step has the potential to introduce failures and to destroy the integrity of the cDNA. Recently, an alternative was introduced, the so called CapFinder approach. It depends on the ability of MMLV reverse transcriptase (RT) to add cytosine residues to the 3′-end of newly synthesised cDNAs upon reaching the 5′-end (cap region) of the mRNA. Usually 2–4 cytosine residues are added, depending on the reaction conditions (4). If an oligonucleotide with oligo(G) or oligo(rG) sequences at its 3′-most end is included in the incubation medium, its terminal 3–4 G residues could base pair with the 2–4 C residues of the newly synthesised cDNA, thus serving as a new template for the RT (template switch). The RT then switches the template and replicates the sequence of the CapFinder oligonucleotide, thus including the complementary CapFinder oligonucleotide sequence at the 3′-end of the newly synthesised cDNA (Fig. 1). This approach allows the amplification of whole cDNA libraries enriched in full-length clones. Enrichment of full-length clones is achieved because the MMLV RT adds C residues preferentially to the cDNA if complete (capped) mRNA serves as template. Variations in the composition of the RT buffer shifts the tendency of the RT to add more C residues to the cDNA end, which makes the entire process more effective (4). Amplification of the cDNA was successfully used for the generation of cDNA libraries and for the production of so-called virtual northern blots, a very useful alternative to mRNA northern blots (3). If these cDNAs are used for 5′-RACE experiments, however, background problems are obvious. Instead of distinct bands, a DNA smear appears if the 5′-RACE product is separated on an agarose gel. This has excluded this promising approach from the important field of 5′-RACE methods. High backgrounds mainly arise from contamination of the 5′-RACE reactions with components of the original CapFinder reaction. The most important contaminants are the CapFinder and oligo(dT) primers. Even residual amounts of these primers result in a high background, because both ideally fit to all cDNAs present in the reaction mixture. To circumvent this problem, strategies to remove all contaminants or to suppress unwanted PCR products are required. Matz and colleagues (5) introduced the PCR suppression effect in combination with CapFinder cDNA synthesis to achieve accurate 5′-RACE products. This refinement of the CapFinder approach seems to work, but, due to the relative complexity of the approach, it has gained insufficient interest. To exclude problems arising from contamination of the primer used for cDNA synthesis, we chose an alternative approach. In our hands, simple, physical removal of all primers is the best approach. To achieve this, we introduced a few, very simple modifications to the cDNA synthesis process. As reported earlier, we introduced a solid phase cDNA synthesis protocol (6) to handle the DNA. MATERIALS AND METHODS RNA (1–10 µg) was isolated from different parts of the locust (Schistocerca gregaria) brain (i.e. the optic lobes, thoracic ganglia or midbrain) or from trophozoites of the human protozoan parasites Entamoeba histolytica and Entamoeba dispar using standard protocols (Trizol Reagent; Life Technologies, Eggenstein, Germany). To isolate the mRNA, a biotinylated oligo(dT) primer [Fig. 2, oligo(dT) I] was bound to avidin-coated polystyrene beads according to the instructions of the manufacturer (Kisker, Mühlhausen, Germany); for each pmol primer, beads with a binding capacity of at least 5 pmol biotin were used. The mRNA is bound according to standard protocols (7) except that it is not eluted from the beads but washed an additional time in 1× cDNA synthesis buffer. For each 1 µg mRNA, 10 pmol oligo(dT) I primer bound to polystyrene beads were used. First strand cDNA synthesis was performed with the mRNA left on the beads using the bound oligo(dT) oligonucleotide as primer. The reaction conditions were as follows: 50 mM Tris–HCl pH 8.3 (45°C), 6 mM MgCl2, 2 mM MnCl2, 1 mM dNTPs, 0.01 M dithiothreitol, 0.2 mg/ml BSA, 10 pM CapFinder primer (CapFinder A or B), 200 U Superscript II (Life Technologies) in 20 µl total volume for 1 h at 45°C. After completion of the reaction the beads were centrifuged, the supernatant removed and the pellet, containing the cDNA bound to the beads, was resuspended in TE buffer. The beads were again centrifuged and the supernatant removed. This procedure was repeated twice, to obtain a washed cDNA preparation that is devoid of any residual primer not incorporated into the cDNA. This preparation could now be used for the 5′-RACE–PCR reaction or stored for several weeks at 4°C (Fig. 1A, step 1). Two alternative strategies could now be chosen. (i) The cDNA could be used for 5′- or 3′-RACE without additional treatment (Fig. 1B, step A). (ii) To enrich for full-length cDNAs, or if only small amounts of material are available, a preamplification with 5′-Primer and 3′-Primer is useful (Fig. 1). Both primers are devoid of the corresponding homopolymeric 3′-regions (Fig. 2), making them ideally suited for effective and specific PCR amplification (3). This amplified DNA has to be captured and bound via its biotin residues to avidin-coated beads to be suitable for subsequent 5′- or 3′-RACE amplification (Fig. 1A, step 4). Aliquots (∼1–2 µl of a 50 µl reaction mix) of this reaction are subjected to a PCR reaction using 5′-Primer and a gene-specific reverse primer (GSRP) derived from the cDNA under investigation (Fig. 1B, step B). 5′-RACE–PCR reactions were either performed under standard conditions (for small cDNAs of <1 kb) or with a LA PCR system (8) for longer cDNAs. Polystyrene beads are far better suited for this purpose compared with magnetic streptavidin beads, because they are smaller and stay without agitation in solution, which is a prerequisite for successful PCR reactions. RESULTS AND DISCUSSION We used this approach to identify 5′-regions of genes isolated in a differential display PCR (DD PCR) (9) screen using different parts of the insect brain (10). In addition, 5′-ends of genes from the protozoan parasites E.histolytica and E.dispar were identified. Among the most problematical tasks in a DD PCR screen is identification of the amplicon 5′-ends. DD PCR makes only small fragments in the most 3′-region of the cDNA accessible, which is a major drawback of this approach. To overcome this, the 5′-RACE has to combine effective amplification with a high reproducibility and a low hands-on time. This is realised in our new DD PCR approach. Both alternative modifications outlined in this study, the direct use of first strand cDNA reactions and the use of PCR preamplified reactions, gave almost identical results. Currently, these methods are in use in our laboratories. One project where the approach is successfully in use is a large DD PCR screen looking for transcripts present in only a single area of the insect brain. We isolated numerous DNAs, each representing a transcript exclusively present in one defined area of the insect brain. To obtain access to the full-length cDNAs we routinely employed our 5′-RACE approach and obtained 5′-regions ranging from a few hundred to several kilobases in length (data not shown). To demonstrate the usefulness of our approach we used known genes of the protozoan parasites E.histolytica and E.dispar. To complete the respective 5′-regions, GSRPs, designed from known regions of the corresponding genes, were used, together with 5′-Primer and the corresponding solid phase CapFinder cDNA. As seen in Figure 3, all reactions gave the anticipated result, yielding a single amplificate of the desired length. The identity of the amplified DNA was revealed by sequencing. In all cases the complete 5′-regions were isolated, including the start codon and a short 5′-untranslated region. The chosen genes are all single copy genes, with some of them transcribed at rather low levels. If we perform these experiments with raw, uncoupled first strand cDNA or amplified cDNA material we obtain a smear of DNA rather than a single DNA population. This holds true even if a second PCR with a nested gene-specific primer is used. The reactions were performed with small aliquots (∼1 µl) of a total reaction (either cDNA or PCR preamplified cDNA) resuspended in a volume of 50 µl. As cDNA synthesis is primed with an oligo(dT) primer rather than with a gene-specific primer, this reaction could be used for the amplification of numerous different 5′-regions. To further evaluate the abilities of this approach, we amplified larger 5′-regions of already identified genes of the human protozoan parasites E.histolytica and E.dispar, which share ∼95% sequence homology. These genes, the cysteine proteinases 1 and 112, are transcribed at relatively low abundance, which is especially true for the second, larger cysteine protease (EhCP112). Alternative 5′-RACE approaches, such as homopolymeric tailing (1), failed to give the desired amplificate. The 5′-RACE reactions were performed with non-amplified first strand cDNA, a gene-specific reverse primer (GSRP; Fig. 1B) and 5′-Primer. These PCRs, which were performed under standard conditions, revealed single DNA bands in the cDNA populations of both species. All of them were of the expected length (∼800 bp for EhCP1 and 1400 bp for EhCP112; Fig. 4). The identities of the amplified DNA populations were verified by sequencing. In addition, the amplified 5′-regions contained the complete 5′-regions including the start codon, which is problematical for amoebae as they usually have very short 5′-untranslated regions. This leads to a loss of the first coding sequences in conventional cDNA libraries made from material of these organisms. Simple and reliable 5′-RACE approaches are therefore very useful to obtain full-length sequences of all cloned genes of these and comparable organisms. Loss of the first few bases of a cDNA during the cDNA library production process is a common problem found in most cDNA libraries. To complete these sequences, very simple and reliable 5′-RACE protocols are required. In addition to its high performance and high efficiency, a single cDNA synthesis is sufficient for numerous 5′-RACE–PCR reactions, reducing hands-on time and use of material to a minimum. The amplification of 1.4 kb transcripts without any specific PCR precautions (without the use of LA PCR systems) demonstrates the usefulness of our novel approach. We were able to obtain even larger parts of unknown 5′-regions using LA PCR reactions (7). Amplifications between 2 and 3 kb were possible in some cases (data not shown). This should significantly facilitate the cloning of larger genes. Our 5′-RACE approach combines numerous advantages compared with other protocols currently in use. It is very effective and yields, in almost every case, the correct amplificate. Usually it is of the desired length, without any contaminating DNA populations. The preference for complete 5′-regions might result from the phenomenon that effective addition of C residues to the cDNA occurs only if complete (capped) mRNA served as the template. The present paper describes a 5′-RACE method that combines simplicity with extraordinary performance. The hands-on time and the chance to introduce experimental errors are reduced to a minimum and optimal results could be obtained in most experiments. The simple solid phase approach opens the potential of the CapFinder approach, which could complement numerous protocols in molecular biology. ACKNOWLEDGEMENTS We would like to thank M. Gewecke for support. This work was funded by grants from the Deutsche Forschungsgemeinschaft to T.R. (DFG Ro 1241) and to I.B. (DFG IB 1744). * To whom correspondence should be addressed. Tel: +49 40 42838 3941; Fax: +49 40 42838 3937; Email: [email protected] View large Download slide View large Download slide Figure 1. Schematic outline of the 5′-RACE approach. (A) cDNA synthesis. The biotin-labelled oligo(dT) primer is bound to avidin-coupled polystyrene-beads. Using these beads, mRNA is isolated and first strand cDNA synthesis starts directly on the beads (step 1). The intrinsic activity of the MMLV RT adds 2–4 C residues to the 3′-end of the newly synthesised cDNA. This region could base pair with the 3′-most part (containing G or rG residues) of the CapFinder A or B primer, which in turn functions as template for the RT. This includes a known sequence complementary to the CapFinder primer sequence at the 3′-end of the newly synthesised cDNA. If only minute amounts of material are available, the cDNA could be preamplified in a LA PCR reaction using 5′-Primer and 3′-Primer (steps 2 and 3). After the PCR reaction the biotinylated product is captured and washed and could now be used in a 5′-RACE–PCR reaction (step 4). (B) 5′-RACE reaction. For the 5′-RACE reaction either non-amplified CapFinder cDNA [see (A) step 1] or PCR amplified cDNA [see (A) step 4] could be used. The 5′-RACE reaction is performed with a gene-specific reverse primer (GSRP) and 5′-Primer, which represents the 5′-most 27 nt of the CapFinder primers. The PCR amplificate represents in both cases the 5′-region of the gene of interest surrounded by the 5′-Primer and gene-specific reverse primer (GSRP). View large Download slide View large Download slide Figure 1. Schematic outline of the 5′-RACE approach. (A) cDNA synthesis. The biotin-labelled oligo(dT) primer is bound to avidin-coupled polystyrene-beads. Using these beads, mRNA is isolated and first strand cDNA synthesis starts directly on the beads (step 1). The intrinsic activity of the MMLV RT adds 2–4 C residues to the 3′-end of the newly synthesised cDNA. This region could base pair with the 3′-most part (containing G or rG residues) of the CapFinder A or B primer, which in turn functions as template for the RT. This includes a known sequence complementary to the CapFinder primer sequence at the 3′-end of the newly synthesised cDNA. If only minute amounts of material are available, the cDNA could be preamplified in a LA PCR reaction using 5′-Primer and 3′-Primer (steps 2 and 3). After the PCR reaction the biotinylated product is captured and washed and could now be used in a 5′-RACE–PCR reaction (step 4). (B) 5′-RACE reaction. For the 5′-RACE reaction either non-amplified CapFinder cDNA [see (A) step 1] or PCR amplified cDNA [see (A) step 4] could be used. The 5′-RACE reaction is performed with a gene-specific reverse primer (GSRP) and 5′-Primer, which represents the 5′-most 27 nt of the CapFinder primers. The PCR amplificate represents in both cases the 5′-region of the gene of interest surrounded by the 5′-Primer and gene-specific reverse primer (GSRP). View largeDownload slide Figure 2. Primers used for the 5′-RACE approach. Sequence of the primers used for the 5′-RACE protocols. Specific primers derived from sequenced genes are not included. mRNA is isolated and cDNA synthesis is primed using oligo(dT) primer I, which contains a biotin residue at its 5′-end. CapFinder primer A or B is included in the first strand synthesis medium to tag the first strand cDNA with a known sequence at its 5′-end. To preamplify the capped cDNA a LA PCR reaction with 5′-Primer and 3′-Primer could be performed; both primers are devoid of the 3′-homopolymeric regions. In the 5′- and 3′-RACE protocols the latter primers are used together with a gene-specific reverse primer (GSRP) and gene-specific sense primers, respectively. View largeDownload slide Figure 2. Primers used for the 5′-RACE approach. Sequence of the primers used for the 5′-RACE protocols. Specific primers derived from sequenced genes are not included. mRNA is isolated and cDNA synthesis is primed using oligo(dT) primer I, which contains a biotin residue at its 5′-end. CapFinder primer A or B is included in the first strand synthesis medium to tag the first strand cDNA with a known sequence at its 5′-end. To preamplify the capped cDNA a LA PCR reaction with 5′-Primer and 3′-Primer could be performed; both primers are devoid of the 3′-homopolymeric regions. In the 5′- and 3′-RACE protocols the latter primers are used together with a gene-specific reverse primer (GSRP) and gene-specific sense primers, respectively. View largeDownload slide Figure 3. 5′-RACE experiments with different genes of the protozoan parasite E.histolytica. Using the 5′-RACE protocol outlined in Figure 1, a number of different 5′-regions of identified genes of the human protozoan parasite E.histolytica were detected. Gene-specific reverse primers (GSRP) were designed from the known sequences of the respective genes and used, together with the 5′-Primer and CapFinder cDNA to amplify the 5′-regions. In lanes 1 and 2 two different 5′-RACE products corresponding to the 5′-ends of cysteine protease cDNAs, in lanes 3 and 4 those of two thioredoxin cDNAs and in lanes 5 and 6 those of two protein disulphide isomerase cDNAs are separated. The smaller band in lane 5 represents a PCR artefact, as it appears even if only the GSRP is present in the PCR reaction. View largeDownload slide Figure 3. 5′-RACE experiments with different genes of the protozoan parasite E.histolytica. Using the 5′-RACE protocol outlined in Figure 1, a number of different 5′-regions of identified genes of the human protozoan parasite E.histolytica were detected. Gene-specific reverse primers (GSRP) were designed from the known sequences of the respective genes and used, together with the 5′-Primer and CapFinder cDNA to amplify the 5′-regions. In lanes 1 and 2 two different 5′-RACE products corresponding to the 5′-ends of cysteine protease cDNAs, in lanes 3 and 4 those of two thioredoxin cDNAs and in lanes 5 and 6 those of two protein disulphide isomerase cDNAs are separated. The smaller band in lane 5 represents a PCR artefact, as it appears even if only the GSRP is present in the PCR reaction. View largeDownload slide Figure 4. 5′-RACE experiments with cysteine proteases of the protozoan parasite E.histolytica. 5′-Regions of the cysteine proteases CP 1 (lanes 1 and 3) and CP 112 (lanes 2 and 4) were amplified from cDNA derived from the protozoan parasites E.histolytica (lanes 1 and 2) and its close relative E.dispar (lanes 3 and 4). Gene-specific reverse primers (GSRP) were chosen that could be anticipated to amplify ∼900 (CP 1, lanes 1 and 3) or 1400 bp (CP 112, lanes 2 and 4). The 5′-RACE amplifications revealed single bands, whose identity was verified by sequencing. View largeDownload slide Figure 4. 5′-RACE experiments with cysteine proteases of the protozoan parasite E.histolytica. 5′-Regions of the cysteine proteases CP 1 (lanes 1 and 3) and CP 112 (lanes 2 and 4) were amplified from cDNA derived from the protozoan parasites E.histolytica (lanes 1 and 2) and its close relative E.dispar (lanes 3 and 4). Gene-specific reverse primers (GSRP) were chosen that could be anticipated to amplify ∼900 (CP 1, lanes 1 and 3) or 1400 bp (CP 112, lanes 2 and 4). The 5′-RACE amplifications revealed single bands, whose identity was verified by sequencing. References 1 Schaefer,B.C. ( 1995) Anal. Biochem. , 227, 255–273. Google Scholar 2 Fromont-Racine,M., Bertrand,E., Pictet,R. and Grange,T. ( 1993) Nucleic Acids Res. , 21, 1683–1684. Google Scholar 3 Franz,O., Bruchhaus,I. and Roeder,T. ( 1999) Nucleic Acids Res. , 27, 1–3. Google Scholar 4 Schmidt,W.M. and Mueller,M.W. ( 1999) Nucleic Acids Res. , 27, e31. 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doi: N/Apmid: N/A

doi: 10.1093/nar/28.22.0pmid: N/A

Abstract This content is only available as a PDF.

Mauffret, O.;Amri, C. El;Santamaria, F.;Tevanian, G.;Rayner, B.;Fermandjian, S.

doi: 10.1093/nar/28.22.4403pmid: 11071926

Abstract Natural and artificial oligonucleotides are capable of assuming many different conformations and functions. Here we present results of an NMR restrained molecular modelling study on the conformational preferences of the modified decanucleotide d(mC1G2mC3G4C5LG6LmC7G8mC9G10)·d(mC11G12mC13G14C15LGL16mC17-G18mC19G20) which contains l deoxynucleotides in its centre. This chimeric DNA was expected to form a right–left–right-handed B-type double-helix (BB*B) at low salt concentration. Actually, it matured into a fully right-handed double helix with its central CLpGL core forming a right-handed Z-DNA helix embedded in a B-DNA matrix (BZ*B). The interplay between base–base and base–sugar stackings within the core and its immediately adjacent residues was found to be critical in ensuring the stabilisation of the right-handed helix. The structure could serve as a model for the design of antisense oligonucleotides resistant to nucleases and capable of hybridising to natural DNAs and RNAs. Received August 25, 2000; Revised and Accepted October 2, 2000. INTRODUCTION l-deoxyribose-containing oligonucleotides could gain particular importance in the context of antisense technology because of their enhanced resistance to nucleases (1,2). However, the pairing between l- and d-DNAs encounters a major steric hindrance and l-DNAs do not, in general, recognise single-stranded natural DNAs or RNAs in either parallel or antiparallel orientations (3). The difficulty of strand association stems mainly from the left-handed nature of homochiral l-DNAs, which does not permit an effective antisense strategy (3,4). Yet, terminally l-modified chimeric oligonucleotides not only exhibit improved hybridisation with natural nucleic acids, but they retain the expected resistance to nucleases (4,5). Also, particularly interesting results are obtained when l-oligonucleotides are designed as ligands for proteins or other small molecules (2,6–8). The conformation of chimeric oligodeoxynucleotides, where l-deoxynucleotide units are inserted into a d-DNA chain, remains rather speculative. An answer to this problem will provide important information on the manner in which these chimeras can recognise RNAs and DNAs to form stable double or triple helices. Until now, the NMR analysis has revealed the loss of stability entailed by the introduction of one or two l-deoxynucleotides within d-DNAs (9). For instance, a punctual incorporation of l-sugars into one strand of a duplex does not impair base pairing and is accompanied by relatively smaller changes in the backbone angles compared to the larger changes on the base stacking. Thus, more substantial modifications in each DNA strand are needed to assess the effect of the chirality on DNA structure. Consequently, we prepared the decanucleotide d(mC1G2mC3G4C5LG6LmC7G8mC9G10)·d(mC11G12mC13G14- C15LGL16mC17G18mC19G20), where the residues of the central CpG step exhibit l-chirality and mC is 5-methylcytosine. The latter substitution was found to facilitate the B→Z transition by generating hydrophobic interactions between the methyl groups and the sugar C1′ and C2′H atoms (10,11). The NMR under conditions of low salt concentration, coupled to a modelling study of this decanucleotide, concludes for a right-handed double-helix (BZ*B, where Z* stands for right-handed Z-DNA), in spite of the fact that a right–left–right double-helix (BB*B, where B* stands for left-handed B-DNA) was expected. Actually, the resulting structure is stabilised by favourable base–base and sugar–base stacking interactions both in the central Z* portion and at the Z*–B junctions. The Tm value of the modified decanucleotide (67.8°C) was barely 5°C below that of its natural counterpart (73.2°C), highlighting its good stability. MATERIALS AND METHODS DNA synthesis Heterochiral oligodeoxynucleotide 5′-d(mCGmCGCLG LmCGmCG) was synthesised using the standard β-cyanoethylphosphoramidite methodology with a Model 381A (Applied Biosystems Inc.) DNA synthesiser (10 µmol scale). Natural 2′-deoxy-d-nucleoside phosphoramidites and derivatised support were from Glen Research. 2′-Deoxy-l-nucleosides were prepared according to slightly modified procedures (3,12). After deprotection with 30% aqueous ammonia for 5 h at 55°C, the crude decanucleotide was purified by anion-exchange chromatography on a DEAE–Sephadex A25 (Pharmacia) column using triethylammonium hydrogen carbonate buffer (pH 7.5, linear gradient from 0.5 to 1.5 M) as eluent. Fractions were analysed by HPLC and those having a purity >95% were combined, co-evaporated with water several times and converted to the sodium salt with 2× Dowex 50 W (yield 15.4 mg). The final purity was >98% as determined by anion-exchange HPLC. MALDI-TOF mass spectrum (3-hydroxypilinic acid/ammonium citrate matrix, Voyager DE, Perspective Biosystems): m/z found 3085.1, calculated 3086.1. UV-melting experiments UV-melting experiments were carried out on a Uvikon 931 (Kontron) spectrometer. The temperature of the cells was controlled by a Huber PD 415 temperature programmer connected to a refrigerated ethylene glycol/water bath (Huber Ministat). Annealing was performed by heating the sample (7.5 µM in 100 mM NaCl, 10 mM cacodylate, pH 7) at 80°C and gradually cooling it to 5°C. Then, the temperature was increased at a rate of 20°C/h and the absorbance measured at 260 nm. Digitised absorbance and temperature values were stored in a computer for subsequent plotting and analysis. NMR spectroscopy NMR spectra of the heterochiral DNA were acquired on a Bruker AMX 500. The spectra were processed with FELIX (Biosym/MSI) software on Silicon Graphics stations. The sample was dissolved in 400 µl of potassium dihydrogen phosphate-disodium hydrogen phosphate buffer containing 1 mM EDTA. The resulting solution consisted of 2 mM in duplex with an ionic strength of 0.1 and a pH of 6.9. Phase sensitive NOESY experiments in TPPI mode (13) in H2O buffer at three different mixing times of 120, 200 and 300 ms were acquired with 4096 complex data points in t2 and 800 real points in t1, with a relaxation delay of 1.5 s, at different temperatures (5–40°C). NOESY spectra in 2H2O buffer were recorded at five different mixing times (60, 120, 180, 240 and 300 ms) with a relaxation delay of 2 s at 20°C. 2048 complex points were acquired in t2 and 800 real points in t1, with a spectral width of 5 kHz. Two-dimensional (2D) P-COSY (14) and 2D TOCSY (15) were used for the proton assignments, 1H–31P experiments (16,17) were used for the phosphorus assignments, and PFG-PEP HSQC (18) for the carbon assignments (19). NMR restraints Interproton distance restraints for structural calculations were obtained from a series of NOESY experiments recorded at various mixing times both in H2O and D2O. The adequate references in both H2O and D2O were taken into account. Distances were classified into four categories (1.8–3, 1.8–4, 1.8–5 and 1.8–6 Å) according to the nOe values and the type of protons (exchangeable or non-exchangeable). Determination of rotamer domains for the backbone torsion angles is made possible using a combination of 1H and 31P-NMR spectroscopy which permits domain definition of α, β, γ, δ, ε and ζ angles as already described (20,21). Molecular dynamics protocol A molecular dynamics simulated annealing protocol driven by experimental restraints was used to solve the solution structure of the molecule. This protocol uses torsion-angle molecular dynamics implementation (22) in the Crystallography and NMR System (CNS) program (23). The two initial strands were used under extended forms, using the CNS protocol generate_extended.inp, to avoid bias of a predetermined form. The refinement procedure was similar to the one used in the original work of Stein et al. (22) with minor modifications. The dynamics protocol was performed in two stages. The first stage involved high temperature molecular dynamics in torsion space. Molecules were equilibrated at 20 000 K, over 60 ps, and cooled slowly at 1000 K over 60 ps. Non-bonded interactions were described by a repulsive quartic term and the calculations were performed under nOe (150 kcal/mol/Å2) and dihedral (5 kcal/mol/degrees2) restraints. The second stage involved Cartesian molecular dynamics consisting first of a cooling step from 1000 to 300 K followed by a 15 ps duration equilibration step at 300 K. During these steps non-bonded interactions were described by Lennard–Jones potentials and electrostatic terms were turned on. At the same time, force constants on dihedral angle restraints were established at 150 kcal/mol/degrees2, while nOe restraints force constants were 150 kcal/mol/Å. 2000 steps of final minimisation were then performed. Coordinates The atomic coordinates of the 10 best structures and the average minimised one have been deposited in the Protein Data Bank with the accession number 1FV7. RESULTS AND DISCUSSION Particular nOes and base pairing observed in water The main nOes observed in H2O are indicated in Figure 1 together with those obtained in 2H2O. NOESY spectra in H2O were recorded primarily to assess the base pairing in the modified decanucleotide through examination of imino and amino protons. Analysis of the fingerprint region of imino protons correlated to amino, aromatic and sugar H1′ protons is presented in Figure 2. The decanucleotide is self-complementary, leading to a duplex with perfect 2-fold symmetry. The H1 imino signals of the eight central G residues are located in the 12.8–13.4 p.p.m. region and reflect the presence of 8 bp, including the two central CL–GL ones. The signals from the two terminal mC–G base pairs, which display significant fraying at 20°C, can be seen only at lower temperatures. The signal at 12.81 p.p.m. is assigned to the guanine H1 imino protons of the two central modified base pairs. Both the chemical shift of these protons and their nOes with the cytosine amino protons (C4H) are fully compatible with a Watson–Crick CL–GL pairing. The H1 imino protons of GL6(16) are further involved in nOes with the sugar H1′ and the base H6 protons of CL5(15) (peaks a and b in Fig. 2). The arrangement of the two l-base pairs is particular. In the standard B-DNA, the interresidue distance GH1 to CH1′/H5 is too large to show such nOe effects and a possible intra-base pair origin is not feasible either, because inside a base pair the mentioned distances are too large to account for the observed nOe intensities. In the amino to aromatic region (not shown), the cross peak at 8.1/7.10 p.p.m. arising from a cytosine is also of great interest. Since inside a cytosine base the distance between the two amino protons and the aromatic proton is >5.3 Å, the observed cross peak could only be assigned to the interstrand nOe C45(15)H42–C415(5)H6, although the two strands are quite identical. The mentioned nOe is incompatible with a B-DNA helix, where the corresponding distance measures 6.5 Å. Thus, NOESY experiments in water provide decisive information relative to the central CLpGL motif in the modified decanucleotide, especially in establishing that this motif maintains a Watson–Crick base pairing while substantially deviating from B-DNA type geometry. Particular nOes and chemical shifts observed in 2H2O The 2D NOESY analysis in the 2H2O solvent confirms the large deviations from B-DNA conformation occurring at the centre of the decanucleotide. First of all, the intrastrand GL6(16)H8–H1′/H2′/H2″/H3′CL5(15) nOes are lacking, and the internucleotide GL6(16)H8–H4′ nOe is unusually strong (Fig. 3). At the same time, the classical B-DNA connectivities are present for the other steps, although the one corresponding to C7(17)H6–G6(16)H1′ appears unusually weak. Secondly, intrastrand intersugar nOes are observed between several protons of the central CLpGL step and its immediately adjacent residues. These include the nOes between H1′ of G4(14) and H2′/H2″/H3′ of CL5(15), and possibly those between H2′/H2″ of G4(14) and H2′/H2″ of CL5(15), which were more difficult to assess due to resonance overlaps. Thirdly, the intranucleotide H2′–H8 and H1′–H8 nOes important for determining the base orientation relative to the sugar are weak and very strong, respectively, suggesting a syn conformation for the GL6(16) base. This is further confirmed by the 13C resonance of the C1′ sugar of GL6(16), shifted 3–4 p.p.m. to lower field, which is typical for syn bases, as underscored particularly by the 13C NMR studies performed on DNA tetraplexes stabilised by guanines in both anti and syn conformations (24). Finally, there are some proton resonances showing abnormally high field shifts. These include the CL5(15) proton resonances H2′ (1.51 p.p.m.), H4′ (3.67 p.p.m. while the other H4′ are located at ∼4.10 p.p.m.), H5″ (3.68 p.p.m.) and, above all, H5′ (2.56 p.p.m., a chemical shift larger by 1.5 p.p.m. compared to that of the other H5′); all of these assignments having been ascertained using 1H–13C HSQC experiments (18,24). For instance, the 13C resonance of the C5′ atom is associated to the H5′ and H5″ proton resonances at 3.6 and 2.55 p.p.m. High field shifts for the cytosine H2′, H5′ and H5″ sugar protons have been seen before in cytosine–guanine Z-steps (25–28). Such effects could be caused by the particular orientation of cytosine in Z-DNA, which places the above mentioned protons directly under the ring current of the neighbouring 5′ and 3′ guanine bases. Structure calculations and analysis The structure of the duplex in solution was resolved by restrained torsion angle molecular dynamics, starting with the two strands in fully extended conformation. Initially, test calculations with synthetic nOes were undertaken with d(CG)5 in order to find the appropriate force constants for the characteristic B-DNA structure. This molecule will be presented in the following section as the reference molecule. Molecular torsion angle dynamics was carried out 50 times using random initial velocities for each structure of the chimeric duplex, attaining the success level of 56% with an overall energy of less than –500 kcal/mol. This high rate was similar to that obtained by Stein et al. (22) for their analysis of B-DNA duplexes. The molecules with the lowest energy were averaged and data for the resulting structure were summarised in Table 1. The r.m.s.d. of the 10 best structures with respect to the average structure was 1.31 Å, seemingly a rather good value, given the conservative restraints used in the system (notably those on nOes), comparable to those previously reported (29). Moreover, the conformation of the four central base pairs of interest, G4CL5GL6C7/G14CL15GL16C17, appeared better resolved (0.60 ± 0.14 Å) relative to the rest of the molecule. Calculated conformation The average structure of the decanucleotide (Fig. 4a) was compared to the structure of the d(CG)5 molecule calculated using synthetic data. Note the Z–Z conformation of the backbone at the central CLpGL step which contrasts with the smooth appearance of the B-DNA backbone in the d(CG)5 molecule. An additional feature is the ‘occupation’ of the major groove at the centre of the modified helix by the two CL–GL central base pairs, which is proper to Z-DNA. The view of the central portion of the molecule, presented in Figure 5, shows that, despite these modifications, the helix is maintained all along. Note, for instance, the reversion of the CL5 sugar relative to the G4 sugar, and that of the GL6 sugar relative to the CL5 sugar; the main direction of the chain, relative to G4, is therefore restored by the second sugar reversion. Also remarkable are the large overlaps occurring between the bases of CL5 and CL15 (interstrand) and the sugar of CL5(15) and the base of GL6(16) (intrastrand), leading to strong stacking interactions. All these features are typical of a Z*-DNA conformation, which accounts for the NMR data, including the particular nOes and the chemical shifts. For example, the position of the H5′ proton of CL5(15) is just above the six-membered ring of GL6(16), which explains its high field shift. Particular stacking interactions in the central portion Figure 6 provides a view of the possible stacking interactions occurring within the two central CL–GL base pairs: (a) B-DNA, (b) Z-DNA, (c) l-modified decanucleotide and (d) the mirror image Z*-DNA. There is no doubt that the stacking pattern of the modified oligomer is clearly of the Z* type. Figure 7 shows the stacking of the central CL–GL base pairs (in red) with their neighbouring mC7–G14 and mC14–G4 base pairs (in green) under different conformations: (a) the B-DNA, (b) the modified decanucleotide, and (c) the mirror-image Z*-DNA. In the modified DNA (b) and in the standard mirror image Z*-DNA (c), the central guanines are pushed away from the centre to the periphery of the helix. The cytosine ring belonging to the neighbouring base pairs (mC7–G14 and mC17–G4) is stacked onto these peripheral guanines. Thus, in both (b) and (c), the central CLpGL step in the Z-DNA structure facilitates the stacking with the immediately preceding and succeeding steps, both of which exhibit some Z-type features. It can be further noted that the stacking of cytosines with guanines at the centre of the helix is better achieved with the l enantiomer than with the d enantiomer. The right-handed BZ*B helix Our results prove, without any ambiguity, that the central CLpGL motif adopts a right-handed Z*-type DNA conformation. This also spans the d-deoxynucleotides that precede and succeed this motif. The originality of the l-modified helix resides in the larger base stacking at its centre compared to the classical left-handed Z-DNA. Indeed, the introduction of an l-deoxynucleotide in a chain of d-deoxynucleotides creates an important structural problem for the double-helix. If the l-sugar points in the same direction as that of the preceding d-sugar, the sense of the helix will be reversed. The major consequence will be a significant loss of base stacking and relative to the major (respectively minor) groove side of the preceding d-deoxyribonucleotide, the l-deoxynucleotide will present its minor (respectively major) groove side. Note that CL5(15) in the modified decanucleotide is constrained to a sugar reversion that helps to maintain base stacking with its preceding d-deoxynucleotide G4(14). The base stacking between the GL6(16) and its succeeding C7(17) is also preserved, but this requires the passage of the guanine in the syn conformation. As a result, the minor and major groove sides of the l-deoxynucleotide bases are not inverted with respect to the l-deoxynucleotide bases. Furthermore, the O4′ atom of the CL5(15) l-sugar is positioned directly above the GL6(16) six-membered ring. Such a system is stabilised via two types of interaction: an intracytidine O4′…H6–C6 hydrogen bond and an n→Π* interaction with the guanidinium ring (30). Within the central CLpGL motif, the guanine prefers to rotate its base and adopt the syn conformation while the cytosine inverts its sugar, which again results in an inversion of the base orientation. At first sight, it may appear intriguing that, in response to stereochemical modifications, the central portion of the molecule adopts a conformation of Z*-DNA type rather than B*-DNA type. Yet, compared with the structural arrangement of the B*-DNA (Fig. 8), the Z*-DNA permits the best stacking at every step, including the steps at the junctions. Our study suggests that a right-handed helix, integrating just two B–Z* junctions, is stable if a good base–base/sugar stacking is achieved. We do not know, however, whether it is the number of l-deoxynucleotides versus the number of d-deoxynucleotides that determines the handedness of the double-helix. We presume that a larger number of l-deoxyribonucleotides would entail the passage to a left-handed double-helix. Biological implications It has been repeatedly demonstrated that DNA fragments constructed from l-nucleotides, despite their better stability towards nucleases, cannot be used as pharmaceutical tools in the antisense strategy (3,4). The main reason invoked was the difficulty for l-DNA strands that resolve into left-handed helical structures to associate to natural DNA or RNA targets. The present results show that l-nucleotides containing DNA chains may assume a right-handed helical structure. This could allow a better strand adjustment to complementary d-DNA or RNA targets, even if good base-pairing requires some accommodation of partners. DNA strands exhibiting BZ*B type conformation could therefore conceivably be used in ‘gene’ therapy (4). Of course, we still need physicochemical and thermodynamical studies of such strands complexed to their natural targets. We believe also that the possible production of DNA-antibodies by Z*-DNA fragments deserves investigation. The N7 atom of the guanine aromatic ring, probably responsible for the production of antibodies against the natural left-handed Z-DNA, is exposed at the periphery of the double-helix in both left-handed Z-DNA and right-handed Z*-DNA and inversion of the helix handedness from left to right could in either case modulate the antigenicity. In addition, single-stranded decanucleotides containing a CpG motif are immunostimulatory (31,32) although the mechanism by which cells detect CpG is not yet totally established. A comparative analysis of a BZ*B strand versus a BB*B strand could offer a good opportunity to test the influence of the CpG chirality on the above-mentioned effect. In conclusion, the CLpGL step at the centre of a l-deoxynucleotide chain may adopt the rare Z-DNA form at physiological salt concentrations. The chimeric molecule apparently resorts to the Z-DNA conformation to solve the problem of stacking continuity along the resulting double-helix. SUPPLEMENTARY MATERIAL Chemicals shifts, the constraints file and the 13C spectra are available as Supplementary Material at NAR Online. ACKNOWLEDGEMENTS We thank M. Agarwal and H. Porumb for critical reading of the manuscript and helpful discussions. * To whom correspondence should be addressed. Tel: +33 1 42 11 49 85; Fax: +33 1 42 11 52 76; Email: [email protected] View largeDownload slide Figure 1. Main nOe connectivities in the central region of the l-modified decanucleotide. Arrows indicate nOes between: the non-exchangeable protons (intra-strand, black); imino and sugar/base protons (intra-strand, green); amino and base protons (inter-strand, red); imino-amino (intra base pair, blue). Also see text. View largeDownload slide Figure 1. Main nOe connectivities in the central region of the l-modified decanucleotide. Arrows indicate nOes between: the non-exchangeable protons (intra-strand, black); imino and sugar/base protons (intra-strand, green); amino and base protons (inter-strand, red); imino-amino (intra base pair, blue). Also see text. View largeDownload slide Figure 2. Expanded NOESY contour plot of the decanucleotide at 20°C in H2O showing the region of connectivities between the imino, amino, aromatic and H1′ resonances. Vertical lines indicate the imino resonances and the names of the linked proton are indicated for each cross peak; (a) and (b) designate the important cross peaks which are discussed in the text. View largeDownload slide Figure 2. Expanded NOESY contour plot of the decanucleotide at 20°C in H2O showing the region of connectivities between the imino, amino, aromatic and H1′ resonances. Vertical lines indicate the imino resonances and the names of the linked proton are indicated for each cross peak; (a) and (b) designate the important cross peaks which are discussed in the text. View largeDownload slide Figure 3. Expanded NOESY contour plot of the decanucleotide at 20°C in D2O showing the aromatic to H1′ region. Sequential connectivities are indicated. The asterisk marks the absent connectivity, which is discussed in the text. View largeDownload slide Figure 3. Expanded NOESY contour plot of the decanucleotide at 20°C in D2O showing the aromatic to H1′ region. Sequential connectivities are indicated. The asterisk marks the absent connectivity, which is discussed in the text. View largeDownload slide Figure 4. Views of (a) the structure of d(CG)5 calculated withthe torsion angle dynamics program using synthetic data (an orange ribbon represents the backbone) and (b) the average structure of the l-modified decanucleotide calculated with experimental data. View largeDownload slide Figure 4. Views of (a) the structure of d(CG)5 calculated withthe torsion angle dynamics program using synthetic data (an orange ribbon represents the backbone) and (b) the average structure of the l-modified decanucleotide calculated with experimental data. View largeDownload slide Figure 5. View of the four central base pairs of the average structure of the l-modified decanucleotide. View largeDownload slide Figure 5. View of the four central base pairs of the average structure of the l-modified decanucleotide. View largeDownload slide Figure 6. Top views showing the stacking interactions of the CpG motif in (a) B-DNA, (b) Z-DNA, (c) l-modified decanucleotide and (d) mirror image Z-DNA. View largeDownload slide Figure 6. Top views showing the stacking interactions of the CpG motif in (a) B-DNA, (b) Z-DNA, (c) l-modified decanucleotide and (d) mirror image Z-DNA. View largeDownload slide Figure 7. Top views of the four central bases in (a) B-DNA, (b) l-modified decanucleotide and (c) mirror image Z-DNA (the central CpG step is in red and the two neighbouring base pairs are in green). View largeDownload slide Figure 7. Top views of the four central bases in (a) B-DNA, (b) l-modified decanucleotide and (c) mirror image Z-DNA (the central CpG step is in red and the two neighbouring base pairs are in green). View largeDownload slide Figure 8. Stacking of one CL–GL base pair (in red) with its preceding G–mC base-pair (in green) in an all B-DNA structure. View largeDownload slide Figure 8. Stacking of one CL–GL base pair (in red) with its preceding G–mC base-pair (in green) in an all B-DNA structure. Table 1. Statistics of the NMR restraints and structures NMR restraints in the complex    Intra-residue distances  168  Inter-residue distances  186  Hydrogen bond restraints  36  Dihedral angle restraints  104  Planarity restraints  10  Total restraints  504  Restraints/residue  25.2  Structure analysis    Average deviations from ideal covalent geometry    Bond length (Å)  0.0017 ± 0.00008  Bond angles (°)  0.364 ± 0.013  Improper angles (°)  0.227 ± 0.004  NOE violations    Number (>0.2 Å)  0  r.m.s.d. of the violations (Å)  0.008 ± 0.001  Dihedral angles violations    Number (>2°)  0  r.m.s.d. of the violations (°)  0.114 ± 0.035  Statistics of superpositions    r.m.s.d. (Å) with respect to the average structure:    10 structures (16 residues)  1.31 ± 0.36  10 structures (eight central residues)  0.60 ± 0.14  NMR restraints in the complex    Intra-residue distances  168  Inter-residue distances  186  Hydrogen bond restraints  36  Dihedral angle restraints  104  Planarity restraints  10  Total restraints  504  Restraints/residue  25.2  Structure analysis    Average deviations from ideal covalent geometry    Bond length (Å)  0.0017 ± 0.00008  Bond angles (°)  0.364 ± 0.013  Improper angles (°)  0.227 ± 0.004  NOE violations    Number (>0.2 Å)  0  r.m.s.d. of the violations (Å)  0.008 ± 0.001  Dihedral angles violations    Number (>2°)  0  r.m.s.d. of the violations (°)  0.114 ± 0.035  Statistics of superpositions    r.m.s.d. (Å) with respect to the average structure:    10 structures (16 residues)  1.31 ± 0.36  10 structures (eight central residues)  0.60 ± 0.14  View Large PDB no. 1FV7. References 1 Urata,H. ( 1999) Yakugaku Zasshi , 119, 689–709. 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Krause, Karen;Wasner, Mark;Reinhard, Wibke;Haugwitz, Ulrike;Dohna, Christine Lange-zu;Mössner, Joachim;Engeland, Kurt

doi: 10.1093/nar/28.22.4410pmid: 11071927

Abstract The tumour suppressor protein p53 has functions in controlling the G1/S and G2/M transitions. Central regulators for progression from G2 to mitosis are B-type cyclins complexed with cdc2 kinase. In mammals two cyclin B proteins are found, cyclin B1 and B2. We show that upon treatment of HepG2 cells with 5-fluorouracil or methotrexate, p53 levels increase while concentrations of cyclin B2 mRNA, measured by RT–PCR with the LightCycler system, are reduced. In DLD-1 colorectal adenocarcinoma cells (DLD-1-tet-off-p53) cyclin B1 and B2 mRNA levels drop after expression of wild-type p53 but not after induction of a DNA binding-deficient mutant of p53. Analysis of the cyclin B2 promoter reveals specific repression of this gene by p53. Transfection of wild-type p53 into SaOS-2 cells shuts off transcription from a cyclin B2 promoter–luciferase construct whereas a p53 mutant protein does not. The cyclin B2 promoter does not contain a consensus p53 binding site. Most of the p53-dependent transcriptional responsiveness resides in its 226 bp core promoter. Taken together with earlier observations on p53-dependent transcription of cyclin B1, our results suggest that one way of regulating G2 arrest may be a reduction in cyclin B levels through p53-dependent transcriptional repression. Received August 16, 2000; Revised and Accepted September 25, 2000. INTRODUCTION The protein p53 is the most important tumour suppressor identified to date. It is mutated in the majority of human tumours, which indicates that its role is important for prevention of malignant transformation (1,2). p53 can influence the cell cycle in several ways. It can cause G1 and G2 growth arrest or apoptosis. The p53 protein plays its part in cell growth regulation and apoptosis induction by engaging in complexes with other proteins or by acting as a transcription factor (1,2). For p53 both function as a transcriptional activator and repressor have been described. Generally, one would expect that genes whose protein products stimulate progression through the cell cycle would be down-regulated by p53. Yet gene expression of inducers of cell cycle arrest or apoptosis should be increased by this tumour suppressor protein (2). The function of p53 as a transcription factor can be modulated by phosphorylation and protein association. Examples are ATM-dependent phosphorylation of p53 at Ser15, regulating the response to DNA damage, and phosphorylation by protein kinases CK2 and Chk2, which are other connectors of p53 with checkpoint controls (3–5). One gene on which p53 acts as an activating transcription factor is bax (6). Induction of bax expression produces a protein with accelerates programmed cell death by counteracting the function of Bcl-2 (7). Another important target for p53-dependent transcriptional activation is the cyclin-dependent kinase inhibitor p21WAF1/CIP1 (8). p53 is thought to exert its function in G1 checkpoint control through p21 (9–11). After induction of transcription of the cell cycle inhibitor by p53, p21 leads to inhibition of cyclin–cdk complexes necessary for the transition from G1 to S phase. However, a function of p21 and p53 in G2 checkpoint control was also observed. It was shown that in colorectal cancer cell lines p53 and p21 are necessary to maintain G2 arrest after γ-irradiation (11). Furthermore, there are some other ways in which p53 can be involved in G2 checkpoint control. In some cell types it has been found that regulation of G2/M progression is contingent upon p53-activated transcription of 14-3-3σ (12). GADD45 transcription is also activated by p53. GADD45 seems to function in controlling the G2/M cell cycle checkpoint induced by ionising but not UV radiation (13). Another notable gene activated by p53 at the transcriptional level is mdm2. The mdm2 protein and its human homologue hdm2 are association partners of p53 and serve to destabilise the tumour suppressor protein (1,2). Regulation by mdm2 is in part based on compartmentalisation through shuttling of this protein between nucleus and cytoplasm (14). p53 function is also connected to the activity of other growth regulatory proteins, like E2F and pRb (1,15,16). These examples illustrate that p53 has many functions. One unexpected finding was its role in DNA repair through its exonuclease activity (17,18). Recently it has been shown that p53-dependent induction of a ribonucleotide reductase is part of the DNA damage checkpoint by causing G2/M arrest and preventing cell death with the help of enzymes required for synthesis of DNA repair precursors (19). An important class of proteins controlling the cell division cycle are the cyclins. They are the regulatory subunits in complexes with cyclin-dependent kinases (cdk). The appearance of cyclins oscillates during the cell cycle (20). Their synthesis is regulated at the transcriptional level and their degradation is controlled by ubiquitin-mediated proteolysis (21). Cyclins are conserved from yeast to man and are involved in controlling cell cycle checkpoints (22,23). An additional level of regulation is provided by cyclin-dependent kinase inhibitors like p21WAF1/CIP1 (20). A central regulator of progression from G2 to mitosis is cyclin B. Cyclin B associates with the cdc2 protein kinase (cdk1) to form maturation-promoting factor (MPF) (24). The MPF complex is essential for transition from G2 to mitosis. In mammalian cells cyclin B exists in two isoforms, cyclins B1 and B2 (25). The activity of cyclin B is regulated by its synthesis, mainly at the transcriptional level, and by its degradation (26). These observations imply a direct role of cyclin B synthesis for MPF kinase activity and ultimately transition from G2 into mitosis. Here we show that cyclin B1 and cyclin B2 mRNA levels decrease upon induction of wild-type p53 and that transcription from the cyclin B2 promoter is repressed by p53. MATERIALS AND METHODS Plasmids The human p21 promoter construct, WWP-luc, and the human p53 expression plasmids, pCMV-p53wt and pCMV-p53mut, were generously provided by Bert Vogelstein (27,28). The reporter construct WWP-luc contains 2.4 kb of the human wild-type WAF1 promoter inserted upstream of a firefly luciferase reporter gene. Both of the p53 expression constructs were in a cytomegalovirus (CMV) promoter-driven expression vector. The plasmid pCMV-p53mut encodes a mutant human p53 protein containing two missense mutations, Pro72→Arg and Val143→Ala. The mouse cyclin B2 promoter constructs were derived from the B2-Luci plasmid which contains a firefly luciferase reporter gene in the pGL3-Basic vector. Some of the B2-Luci constructs have been described by us (29). New constructs were prepared by PCR by using the following primers together with the respective antisense oligonucleotides: Mut-930, 5′-CTGATGGGGTAGCCTACGCTCAAGT-3′; Mut-845, 5′-TTTGTTTTGATCGATCATTTTTGTTTTCTGTCTTGTC-3′; Mut-750, 5′-CCGTCATTTGGTAGGTAGTTTCT-3′; Mut-460, 5′-CCCAGAGACCACTTTTAAAGACATATGTC-3′; Mut-305, 5′-GAAAATAACCGGGTGTACAAGGAAACA-3′; CDE-Mut, 5′-CAATAGTGCGTCAGCATTACGGTATTTGAATCGCGGACCGG-3′; CHR-Mut, 5′-CAGCGGCGCGGTATGCATATCGCGGACCGGGCGGTGG-3′. Mutations are given in italics. Cyclin B2 promoter deletion constructs were created by employing the following upstream primers: –684B2, 5′-GGGGTACCGCACATCACACCGTCATTTG-3′; –453B2, 5′-GGGGTACCGAGGAAGTAAGGTCAGAAGTAG-3′; –226B2, 5′-GGGGTACCGCTATGACAAGCAAATACAAGC-3′. The plasmid pRL-null (Promega) contains a cDNA encoding Renilla luciferase. All of the construct DNAs were purified through anion exchange columns (Qiagen) and confirmed by restriction analysis and sequencing. Cell culture and chemotherapeutics HepG2, Hep3B and SaOS-2 cells were obtained from DSMZ (Braunschweig, Germany) and cultured in a humidified atmosphere with 5% CO2 at 37°C. HepG2 cells were grown in RPMI 1640 medium (Biochrom) supplemented with 10% foetal calf serum (FCS) (Biochrom). Hep3B cells were maintained in medium containing Minimum Essential Medium with Eagle’s salts (MEM Eagle; Biochrom), 0.1 mM non-essential amino acids and 10% FCS. Cells were treated with chemotherapeutic agents for 24 h at concentrations of 2.5 and 25 µg/ml 5-fluorouracil (5-FU) and 10 and 100 µg/ml methotrexate (MTX). The concentrations of chemotherapeutics were at levels derived from cancer treatment protocols which were employed in previous experiments (30). SaOS-2 cells were cultured in McCoy’s 5A modified medium (Biochrom) supplemented with 15% FCS. Inducible cell lines D.P53 A2 and 175 A4 were kindly provided by Bert Vogelstein. Both cell lines are derivatives of the colorectal carcinoma cell line DLD-1, which has endogenous mutant p53 alleles (31). They were grown in 10% FCS in McCoy’s 5A modified medium containing 400 µg/ml geneticin (Gibco BRL) and 250 µg/ml hygromycin (Roche). Expression of p53 wt or p53R175H, respectively, is regulated by a modified tetracycline (tet)-regulated gene expression system (tet-off system) (31,32). p53 expression was kept repressed in the presence of 20 ng/ml doxycycline (Sigma) but was induced upon removal of doxycycline from the culture medium. For induction of expression of wild-type or mutant p53, cells were washed three times with phosphate-buffered saline (PBS) and placed in medium lacking doxycycline. Fluorescence-activated cell sorting (FACS) analysis Cells were harvested, washed twice in PBS/EDTA (1 mM) and fixed with 75% ethanol in PBS/EDTA for at least 12 h at 4°C. Cells were centrifuged and resuspended in 1 ml PBS/EDTA containing 50 µg/ml RNase A (Sigma). Cells were stained with propidium iodide (Sigma) at a final concentration of 60 µg/ml and filtered through a 35 µm pore size cell strainer (Falcon). Flow cytometry was performed by using a FACSCalibur sorter (Becton Dickinson). A total of 10 000–20 000 cells were analysed with the CELLQuest software (Becton Dickinson). Transfections and luciferase assays SaOS-2 cells were transfected by lipofection with Fugene 6 (Roche) according to the manufacturer’s instructions. Transfections were done in triplicate. Exponentially growing cells were plated at a density of 5 × 104/well in 0.5 ml medium in 24-well plates. Cells were cultured overnight before transfection. Unless otherwise indicated, 125 ng of luciferase reporter constructs were co-transfected with 25 ng of constructs expressing wild-type or mutant p53 proteins and 25 ng of Renillaluciferase expression vector (pRL-null; Promega) as an internal control. DNA amount was held constant in all experiments by adjusting with pcDNA3.1/His C (Invitrogen). The quality and quantity of several independent DNA preparations were checked photometrically and visually on agarose gels. The cells were harvested 24 h after transfection by lysis with passive lysis buffer (Promega). Firefly and Renilla luciferase activities were assayed with the Dual Luciferase Assay System (Promega) as suggested by the manufacturer. The firefly luciferase activity was normalised to Renilla luciferase activity to compensate for variability in transfection efficiencies. Western blot analysis Cell pellets were lysed in RIPA lysis buffer (33) containing a cocktail of protease inhibitors (Complete; Roche). The protein concentration of each cell lysate was determined with the Bio-Rad protein assay kit (Bio-Rad). Ten micrograms of total protein were separated in a 10% SDS–polyacrylamide gel and transferred to a polyvinylidene difluoride transfer membrane (Hybond-P; Amersham Pharmacia Biotech) according to standard procedures (33). Western blotting using a 1:1000 dilution of the anti-p53 mouse monoclonal antibody DO-1 (Calbiochem) was performed and analysed with an ECL Western blotting analysis system (Amersham Pharmacia Biotech) according to the manufacturer’s instructions. RNA extraction and LightCycler RT–PCR analysis Total RNA was extracted from 5 × 106 cells using RNeasy kits (Qiagen) as described by the manufacturer and quantified by optical density. One-step RT–PCR was performed with a LightCycler instrument (Roche) in a total volume of 20 µl containing 50 ng of total RNA, 4 mM MgCl2, 0.5 µM each primer, LightCycler RT–PCR Reaction Mix SYBR Green I (1×) and LightCycler RT–PCR Enzyme Mix (Roche). The protocol consists of four programs: reverse transcription of template RNA, denaturation of the cDNA/RNA hybrid, amplification of cDNA and melting curve analysis for product identification. Reverse transcription was performed at 55°C for 10 min. The denaturation and amplification conditions were 95°C for 30 s followed by up to 34 cycles of PCR. Each cycle of PCR included immediate denaturation at 95°C, 10 s of primer annealing at 55°C and 15 s of extension/synthesis at 72°C. The temperature ramp was 20°C/s, except when heating to 72°C, when it was 2°C/s. At the end of the extension step fluorescence of each sample was measured to allow quantification of the RNA. After amplification a melting curve was obtained by heating at 20°C/s to 95°C, cooling at 20°C/s to 65°C and slowly heating at 0.1°C/s to 95°C with fluorescence data collection at 0.1 °C intervals. The following primers were used for PCR: B1-hum-for, 5′-AAGAGCTTTAAACTTTGGTCTGGG-3′; B1-hum-rev, 5′-CTTTGTAAGTCCTTGATTTACCATG-3′; B2-hum-for, 5′-AAAGTTGGCTCCAAAGGGTCCTT-3′; B2-hum-rev, 5′-GAAACTGGCTGAACCTGTAAAAAT-3′; GAPDH-for, 5′-CAGTCCATGCCATCACTGCC-ACCCAG-3′; GAPDH-rev, 5′-CAGTGTAGCCCAGGATGCCCTTGAG-3′. These primer pairs result in PCR products of 317 (cyclin B1) (34), 351 (cyclin B2; GenBank accession no. AL080146) and 303 bp (GAPDH) (35). Quantitative analysis of the LightCycler data was performed employing LightCycler analysis software. The data analysis is divided into two parts: specificity control of the amplification reaction using the melting curve program of the LightCycler software, followed by use of the quantification program. The SYBR Green I signal of each sample is plotted versus the number of cycles. Using the LightCycler analysis software background fluorescence is removed by setting a noise band. This fluorescence threshold is used to determine cycle numbers that correlate inversely with the log of the initial template concentration. To this end the log-linear portions of the amplification curves are identified and best fit lines calculated. The crossing points (CP) are the intersections between the best fit lines of the log-linear region and the noise band. These crossing points correlate inversely with the log of the initial template concentration (LightCycler Operator’s Manual, Version 3.0, May 1999, Roche). The CP determined for cyclin B2 mRNA were normalised to those of GAPDH to compensate for variability in RNA amount and for exclusion of general transcriptional effects. We calculated fold reduction (FR) since our experiments yielded a repressive effect: FR = 2(CP1–CP2). Fold induction can be calculated by the same formula by reversing the signs. CP1 indicates the crossing point of an RNA sample from cells treated with chemotherapeutic agents; CP2 indicates the crossing point of an RNA sample originating from untreated cells. The mRNA levels of untreated cells were set at 100%. Remaining mRNA levels after treatment with chemotherapeutics were estimated using calculated fold reductions and given as calculated mRNA% in the figures relative to levels of untreated samples. RESULTS Chemotherapeutics reduce expression of cyclin B2 in p53-positive cells Treatment of cells with chemotherapeutics like 5-FU and MTX has been shown to induce expression of p53 (30,36). The two hepatocellular carcinoma cell lines HepG2 and Hep3B were treated with increasing amounts of the two agents and analysed for expression of p53 protein. In HepG2 cells we find a clear increase in p53 protein over the basal expression level detected in untreated cells after addition of 5-FU or MTX. In both cases protein expression increases further with larger amounts of agent (Fig 1). Hep3B cells serve as negative controls since they lack intact p53 genes (30). In these cells no expression of p53 protein is observed (Fig. 1). In p53-positive HepG2 cells we looked at expression of cyclin B2 mRNA. The quantification of low abundance cellular transcripts requires sensitive techniques. With a method to determine mRNA expression employing the LightCycler system we compared mRNA levels in the same samples as in Figure 1. This method is based on the analysis of RT–PCR products. Continuous fluorescence detection of amplifying cDNA allows rapid and accurate quantification of initial transcript amount. A simple and general method for monitoring product synthesis with the double-stranded DNA dye SYBR Green I provides an estimation of initial template amounts and with that comparison of mRNA levels in different samples. cyclin B2 mRNA levels were normalised to GAPDH expression. Results are reported as calculated mRNA%. (For a detailed description of the calculations see Materials and Methods.) We compared cyclin B2 mRNA levels from untreated HepG2 cells with two concentrations of MTX or 5-FU. In both experiments the expression of cyclin B2 mRNA was clearly down-regulated. The down-regulation appears to be concentration-dependent since higher concentrations of cancer therapeutics lead to lower cyclin B2 mRNA levels (Fig. 2). A specific increase in p53 expression results in down-regulation of cyclin B1 and B2 mRNA levels After finding that an increase in p53 protein is observed together with a decrease in cyclin B2 mRNA upon chemotherapeutics treatment of cells, we were interested in examining how specific this observation is to the up-regulation of p53. In order to look specifically at the influence of p53 we employed the DLD-1 colorectal adenocarcinoma cell line, which has a p53 mutant background. These cells were stably transfected with a system that permits tet-off regulation for a wild-type or a DNA-binding mutant of p53 (p53R175H) (31). This system allows for selective induction of p53 wild-type or mutant protein in the two created cell lines by removal of doxycycline from the medium (31,32). p53 protein is already induced after 3 h and continues to increase for the next 6 h after culturing the cells in medium lacking doxycycline (Fig. 3A). In the same cultures we tested for cyclin B1 and B2 mRNA expression. Upon expression of wild-type p53 both cyclin mRNAs were reduced after 3 h and dropped further at 6 h, reaching residual levels at 9 h of 1.1 and 2.4% for cyclin B1 and B2, respectively (Fig. 3B). In the control experiment with induction of the p53 DNA-binding mutant expression of both cyclins increased slightly over mRNA levels without p53 induction (Fig. 3B). We also checked cells from this experiment by flow cytometry for the appearance of a sub-G1 DNA staining population. Induction of wild-type p53 expression leaves the portion of sub-G1 staining cells similarly low for up to 6 or 9 h while the number of cells in G2 had increased significantly at 6 h. For the 12 and 24 h time points a strong increase in the sub-G1 population was observed (Fig. 3C). Control experiments with mutant p53 induction did not show any significant change in the DNA content of cells compared to uninduced cells for the monitored 24 h period after mutant p53 expression (data not shown). The promoter activity of cyclin B2 is down-regulated by p53 We analysed whether expression of p53 has an influence on the cyclin B2 promoter in SaOS-2 cells with co-transfections of p53-expressing plasmids and cyclin B2 promoter–luciferase constructs. As a control we first tested whether wild-type p53 but not a mutant of this protein could activate transcription from a p21Waf1/Cip1 promoter (Fig. 4A). The p53 mutant, which contains a disabled DNA-binding domain, does not activate transcription significantly. The luciferase values for this control were the same as the background from the pGL3 luciferase vector lacking any promoter (data not shown). However, wild-type p53 triggered a strong increase in transcription from the p21 promoter of >12-fold over the negative p53 mutant (Fig. 4A). Analysis of the influence of p53 on the cyclin B2 promoter–luciferase constructs shows an opposite effect. Increasing amounts of wild-type p53-expressing plasmid reduce expression from the cyclin B2 promoter down to background levels. With the largest amount of wild-type p53 plasmid co-transfected the observed repression in this assay was ∼30-fold (Fig. 4B and C). It appears from a number of experiments that cyclin B2 expression is completely shut down, to below the detection limit (data not shown). The calculated factor for repression is dependent on the basal expression level and therefore might be even higher. The cyclin B2 promoter might be completely shut off upon expression of wild-type p53. Repression of cyclin B2 expression by p53 does not employ a p53 consensus binding site and resides in the 3′-part of the promoter The next question we wished to answer was through which DNA element in the cyclin B2 promoter is this strong repression made possible? A binding consensus for p53 has been published (37). We inspected the cyclin B2 promoter for a site that would match the p53 consensus and did not find any element that could serve as a p53 binding site. We then started a first preliminary search for the DNA element through which p53 represses cyclin B2 transcription. We compared the mouse and human cyclin B2 promoter sequences and found a number of sites which are conserved in both promoters (M.Wasner, unpublished data). Among these sites there are three NF-Y-binding CCAAT boxes, the cell cycle-dependent element (CDE) and the cell cycle genes homology region (CHR) (26,29). We individually mutated nine of these homology regions on the basis of the mouse cyclin B2 promoter–luciferase construct (Fig. 5A). We analysed these constructs for repression by co-expressed wild-type p53 versus mutant p53. All mutant cyclin B2 promoter constructs were still strongly down-regulated by wild-type p53 (Fig. 5B). A significant change in the repression factor compared to the wild-type construct is only seen with mutant promoters which display a low total activity. Since this search did not reveal the dominant site for repression by p53 we constructed three truncation mutants of the cyclin B2 promoter (Fig. 5A). For the shortest mutant repression drops to 18-fold, compared to 29-fold for the wild-type construct (Fig. 5B). Therefore, nearly all of the p53-dependent repression still resides even in the shortest truncation mutant with 226 bp remaining from the translational start at the 3′-end of the cyclin B2 promoter. DISCUSSION p53 activates or represses transcription of a number of central regulators of cell division and apoptosis. We find that cyclin B2 transcription is shut down by expression of wild-type p53. The other B-type cyclin, cyclin B1, has already been shown to be down-regulated by p53 at the transcriptional level (38,39). Therefore, both mammalian cyclin B genes are repressed by p53. One way of increasing p53 levels is by incubating cells with chemotherapeutic agents. Treatment of cells with chemotherapeutics certainly leads to profound changes in the expression of many genes. Therefore, even though we see an up-regulation of p53 protein and a down-regulation of cyclin B2 mRNA these two observations might not be directly connected (Figs 1 and 2). Consequently, in later experiments we looked at colorectal carcinoma cells with a p53 mutant background in which wild-type or mutant p53 could be selectively induced. We observed an effective decrease in both cyclin B1 and B2 mRNA levels upon expression of wild-type but not mutant p53 (Fig. 3). This reduction is already seen 3 h after removal of antibiotic from the DLD-1-tet-off-p53 cells (Fig. 3B). Considering the time needed for expression of p53 and degradation of cyclin B mRNA this may implicate an immediate response by down-regulation of cyclin B transcription. General effects like the induction of apoptosis seem to appear later, since a significant sub-G1 cell population, as judged by DNA staining and FACS analysis, is not observed until 9 h after induction of p53. However, at 6 h an increase in the G2/M cell population can already be detected (Fig. 3C). These observations are consistent with earlier results when p53-dependent transcriptional repression and cell cycle control were analysed. It has been reported that high levels of p53 in TR9-7 skin fibroblasts lead to a reduction in cdc2 and cyclin B1 mRNA levels and to G2 cell cycle arrest (39). Another study showed that overexpression of cyclin B1 can overcome p53-dependent G2 arrest in an ovarian cancer cell line (38). Taken together, the results from these two recent papers and our experiments indicate that the timing of the decrease in cyclin B mRNAs precedes G2 arrest and appearance of apoptotic cells (Fig. 3). In order to obtain more information on how cyclin B2 is down-regulated we performed co-transfections of p53 wild-type and mutant constructs and analysed their effect on expression of a cyclin B2 promoter–luciferase reporter. Cyclin B2 expression is repressed down to background levels upon co-transfection of wild-type p53 in a concentration-dependent manner (Fig. 4). In previous reports on the cyclin B1 promoter reduction of its transcription by p53 was not as pronounced as described here for cyclin B2 (38,39). However, when a similar experimental approach was taken, e.g. co-transfection of a reporter and wild-type p53 in SaOS-2 cells, repression of the cyclin B1 promoter was close to vector background levels (38). In summary, these results show that concentrations of both B-type cyclins are substantially reduced when p53 is up-regulated. Human and mouse cyclin B2 transcription are both repressed by p53, but their promoters do not display a p53 consensus binding site. In initial experiments to elucidate by which elements p53 can repress the cyclin B2 promoter we analysed regions homologous in the human and mouse promoters. We mutated all homologous regions on the basis of the mouse construct and tested if expression from these plasmids was still repressed by p53 (Fig. 5). Among these promoter constructs was the Y-1,2,3-Mut plasmid in which three CCAAT boxes had been mutated. It has been shown that p53 can repress the hsp70 promoter through interaction with the CCAAT box-binding factor (CBF or NF-Y) and that an NF-Y-binding CCAAT box in the cdc2 promoter is the site for repression by p53 (40–42). We found earlier that the three CCAAT boxes in the 3′-region of the cyclin B2 promoter are the main activators for transcription (29). Now we observe that transcription from the Y-1,2,3-Mut plasmid is still repressed by wild-type p53 but not mutant p53 (Fig. 5B). However, there is a change in the factor by which p53 can repress Y-1,2,3-Mut. Yet, as with the Mut-845 construct, Y-1,2,3-Mut also shows a low absolute activity. Therefore, repression by p53 down to background simulates a lower repression factor in these cases (Fold reduction, Fig. 5B). Absolute activity of the CHR mutant is increased over the wild-type promoter. The CHR element is responsible for cell cycle-dependent repression of cyclin B2. Destruction of this site leads to derepression of this promoter which yields a higher average activity in an asynchronous growing cell population (Fig. 5B and data not shown). Since the constructs created on the basis of homology between mouse and human promoters did not yield a mutant in which repression was lost to a significant extent we tested three large deletion mutants of the cyclin B2 promoter. The three mutants retained the ability for p53-dependent down-regulation (Fig. 5). The fact that even the –226B2 mutant is still repressed similarly to the full-length wild-type promoter suggests that this promoter segment is sufficient to mediate p53-dependent repression. The three CCAAT boxes responsible for most of the activation and the CDE/CHR cell cycle elements are found in this most proximal segment (29). Although this core promoter contains most of the sites essential for the properties of the cyclin B2 promoter we could show that these elements are not the sites through which p53 down-regulates cyclin B2 transcription (Fig. 5). It has been suggested that repression by p53 can be regulated through TATA boxes (43). However, many of the promoters of cell cycle genes, including the cyclin B2 promoter described here, do not contain TATA elements for transcriptional initiation. Therefore, further experiments are required to elucidate the exact mechanism of repression. Cyclin B is essential for cdc2 kinase to form MPF, which in turn induces transition from G2 to mitosis. Human cyclins B1 and B2 are both able to serve as the activating partner for cdc2 kinase, with peak activity at mitosis (44). Innocente et al. recently showed that histone kinase activity in cdc2 immunoprecipitates and in co-immunoprecipitates with cyclins B1 and B2 is reduced upon induction of p53. This reduction in activity is not due to reduced expression of cdc2 kinase or a change in its tyrosine phosphorylation. However, there is a marked decrease in cyclin B1 mRNA and protein (38). We now find that cyclin B2 is down-regulated after increased expression of p53. It is obvious that expression of both cyclin B1 and B2 have to be reduced to ultimately shut off cdc2 kinase activity and slow down execution of G2/M progression. This is corroborated by the fact that re-expression of cyclin B1 alone can overcome p53-induced G2 cell cycle arrest (38). Taken together, these observations indicate that transcriptional repression of cyclin B by p53 is important for cell cycle progression (Fig. 6). Other factors transcriptionally induced by p53 have been implicated in G2/M checkpoint control. One example is 14-3-3σ, which contains a p53-responsive element in its promoter and is able to control G2 arrest by employing compartmentalisation through cytoplasmic retention of cyclin B/cdc2 (12,45). There are multiple ways by which p53 may be involved in the different checkpoints. Cell type-specific differences exist. Some of the pathways are redundant and some will have to cooperate to exert the p53 checkpoint function (1,2,18,46). By different means p53 seems to play an important role in reducing the concentration of active cyclin B–cdc2 complex in the nucleus to maintain G2 arrest. Repression of cyclin B transcription by p53 is part of this regulatory network. ACKNOWLEDGEMENTS We wish to thank Jana Lorenz for technical assistance and Bert Vogelstein for generously providing the DLD-1 tet-off cells in addition to the p21 promoter and p53 expression plasmids. K.E. is supported by grants from the Bundesministerium für Bildung und Forschung (through the IZKF) and the Deutsche Forschungsgemeinschaft. * To whom correspondence should be addressed. Tel: +49 341 972 5900; Fax: +49 341 971 2209; Email: [email protected] View largeDownload slide Figure 1. Increase in p53 protein after treatment with chemotherapeutics. p53-positive HepG2 and p53-negative Hep3B cells were treated with 5-FU or MTX and analysed for p53 expression by western blotting. Untreated cells served as a control. HepG2 and Hep3B cells were treated with 5-FU or MTX, respectively, as indicated. View largeDownload slide Figure 1. Increase in p53 protein after treatment with chemotherapeutics. p53-positive HepG2 and p53-negative Hep3B cells were treated with 5-FU or MTX and analysed for p53 expression by western blotting. Untreated cells served as a control. HepG2 and Hep3B cells were treated with 5-FU or MTX, respectively, as indicated. View largeDownload slide Figure 2.cyclin B2 mRNA levels decrease upon treatment with chemotherapeutics in p53-positive cells. The same samples from HepG2 cells as used in Figure 1 were analysed with the LightCycler system for their change in cyclin B2 mRNA concentration. Expression levels were normalised to GAPDH mRNA levels. For a detailed description of calculations see Materials and Methods. View largeDownload slide Figure 2.cyclin B2 mRNA levels decrease upon treatment with chemotherapeutics in p53-positive cells. The same samples from HepG2 cells as used in Figure 1 were analysed with the LightCycler system for their change in cyclin B2 mRNA concentration. Expression levels were normalised to GAPDH mRNA levels. For a detailed description of calculations see Materials and Methods. View large Download slide View large Download slide View large Download slide Figure 3. cyclin B1 and cyclin B2 mRNA levels from endogenous genes are down-regulated upon selective induction of p53. DLD-1 colorectal cancer cells with a tet-off-regulated gene for wild-type or a DNA-binding mutant of p53 were induced to increase expression of p53 or mutant p53 by removal of doxycycline. (A) Western analysis of p53 induction. Lanes 1 and 5 represent control lysates from DLD-1-tet-off-p53-wt and DLD-1-tet-off-p53-mut cells in the presence of doxycycline, respectively. In lanes 2 and 6, 3 and 7 and 4 and 8 p53 induction 3, 6 and 9 h after antibiotic removal, respectively, is shown. (B) Relative mRNA levels for cyclin B1 and B2 measured by LightCycler RT–PCR. Levels were standardised to expression of GAPDH and mRNA concentrations from control DLD-1-tet-off cells in medium containing doxycycline were set as the 100% reference. (C) DLD-1-tet-off-p53-wt cells were harvested before removal of doxycycline from the medium as a control or at indicated times after elimination of doxycycline from the medium. The cells were stained with propidium iodide and subjected to flow cytometry. DNA staining with propidium iodide versus FL2 pulse width is depicted. View large Download slide View large Download slide View large Download slide Figure 3. cyclin B1 and cyclin B2 mRNA levels from endogenous genes are down-regulated upon selective induction of p53. DLD-1 colorectal cancer cells with a tet-off-regulated gene for wild-type or a DNA-binding mutant of p53 were induced to increase expression of p53 or mutant p53 by removal of doxycycline. (A) Western analysis of p53 induction. Lanes 1 and 5 represent control lysates from DLD-1-tet-off-p53-wt and DLD-1-tet-off-p53-mut cells in the presence of doxycycline, respectively. In lanes 2 and 6, 3 and 7 and 4 and 8 p53 induction 3, 6 and 9 h after antibiotic removal, respectively, is shown. (B) Relative mRNA levels for cyclin B1 and B2 measured by LightCycler RT–PCR. Levels were standardised to expression of GAPDH and mRNA concentrations from control DLD-1-tet-off cells in medium containing doxycycline were set as the 100% reference. (C) DLD-1-tet-off-p53-wt cells were harvested before removal of doxycycline from the medium as a control or at indicated times after elimination of doxycycline from the medium. The cells were stained with propidium iodide and subjected to flow cytometry. DNA staining with propidium iodide versus FL2 pulse width is depicted. View largeDownload slide Figure 4. The cyclin B2 promoter is down-regulated by p53. Luciferase reporter constructs and p53 wild-type and mutant expression plasmids were co-transfected into SaOS-2 cells. All experiments were standardised to Renilla luciferase activity expressed from the co-transfected pRL-null vector. The total amount of transfected DNA was held constant. (A) Stimulation of transcription from a p21 reporter construct as positive control. An aliquot of 125 ng of the B2-Luci reporter plasmid and 25 ng of plasmids expressing wild-type p53 or a DNA binding-deficient p53 mutant were transfected. The expression level with co-transfection of mutant p53 was used as the 100% reference. (B) The cyclin B2 reporter plasmid was co-transfected with 25 ng of vector control and increasing amounts of the plasmid expressing wild-type p53. The control served as the 100% standard. (C) Western blot depicting p53 protein after co-transfection of increasing amounts of a p53-expressing plasmid. Lanes represent lysates from the experiments shown in (B). Lane 1 is without transfection of p53 plasmid; lanes 2–4 are lysates from cells transfected with 1, 5 and 25 ng of wild-type p53-expressing plasmid, respectively. View largeDownload slide Figure 4. The cyclin B2 promoter is down-regulated by p53. Luciferase reporter constructs and p53 wild-type and mutant expression plasmids were co-transfected into SaOS-2 cells. All experiments were standardised to Renilla luciferase activity expressed from the co-transfected pRL-null vector. The total amount of transfected DNA was held constant. (A) Stimulation of transcription from a p21 reporter construct as positive control. An aliquot of 125 ng of the B2-Luci reporter plasmid and 25 ng of plasmids expressing wild-type p53 or a DNA binding-deficient p53 mutant were transfected. The expression level with co-transfection of mutant p53 was used as the 100% reference. (B) The cyclin B2 reporter plasmid was co-transfected with 25 ng of vector control and increasing amounts of the plasmid expressing wild-type p53. The control served as the 100% standard. (C) Western blot depicting p53 protein after co-transfection of increasing amounts of a p53-expressing plasmid. Lanes represent lysates from the experiments shown in (B). Lane 1 is without transfection of p53 plasmid; lanes 2–4 are lysates from cells transfected with 1, 5 and 25 ng of wild-type p53-expressing plasmid, respectively. View large Download slide View large Download slide Figure 5. Repression of cyclin B2 transcription by p53 is not mediated by regions homologous in the mouse and human promoters and resides just upstream of the coding region. (A) Anatomy of the cyclin B2 promoter. Regions homologous in the human and mouse promoter are indicated by their names or as their respective mutants with numbers for their location counted in nucleotides from the first codon. For the three truncation mutants the remaining elements are indicated. (B) Analyses of the different mutant reporters upon transfection of wild-type or mutant p53. Experimental conditions were the same as in Figure 4. The fold reduction was calculated by dividing luciferase activities from p53 mutant transfections by values from p53 wild-type transfections. Fold reductions shown in parentheses were calculated for reporters with low general activities which are closer to background levels and therefore result in lower repression factors. View large Download slide View large Download slide Figure 5. Repression of cyclin B2 transcription by p53 is not mediated by regions homologous in the mouse and human promoters and resides just upstream of the coding region. (A) Anatomy of the cyclin B2 promoter. Regions homologous in the human and mouse promoter are indicated by their names or as their respective mutants with numbers for their location counted in nucleotides from the first codon. For the three truncation mutants the remaining elements are indicated. (B) Analyses of the different mutant reporters upon transfection of wild-type or mutant p53. Experimental conditions were the same as in Figure 4. The fold reduction was calculated by dividing luciferase activities from p53 mutant transfections by values from p53 wild-type transfections. Fold reductions shown in parentheses were calculated for reporters with low general activities which are closer to background levels and therefore result in lower repression factors. View largeDownload slide Figure 6. An increase in p53 may influence the transition from G2 to mitosis by repressing transcription of cyclin B. Repression of cyclin B1 and B2 transcription by p53 results in lower cyclin B protein concentrations. Reduced amounts of cyclin B leave the cdc2 kinase inactive, which may contribute to an arrest of the cell cycle in G2. View largeDownload slide Figure 6. An increase in p53 may influence the transition from G2 to mitosis by repressing transcription of cyclin B. Repression of cyclin B1 and B2 transcription by p53 results in lower cyclin B protein concentrations. Reduced amounts of cyclin B leave the cdc2 kinase inactive, which may contribute to an arrest of the cell cycle in G2. References 1 Levine,A.J. ( 1997) Cell , 88, 323–331. 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Pitt, Melinda;Gallegos, María-Trinidad;Buck, Martin

doi: 10.1093/nar/28.22.4419pmid: 11071928

Abstract Transcription initiation by the σ54 RNA polymerase requires specialised activators and their associated nucleoside triphosphate hydrolysis. To explore the roles of σ54 in initiation we used random mutagenesis of rpoN and an in vivo activity screen to isolate functionally altered σ54 proteins. Five defective mutants, each with a different single amino acid substitution, were obtained. Three failed in transcription after forming a closed complex. One such mutant mapped to regulatory Region I of σ54, the other two to Region III. The Region I mutant allowed transcription independently of activator and showed reduced activator-dependent σ54 isomerisation. The two Region III mutants displayed altered behaviour in a σ54 isomerisation assay and one failed to stably bind early melted DNA as the holoenzyme; they may contribute to a communication pathway linking changes in σ to open complex formation. Two further Region III mutants showed gross defects in overall DNA binding. For one, sufficient residual DNA binding activity remained to allow us to demonstrate that other activities were largely unaffected. Changes in DNA binding preferences and core polymerase-dependent properties were evident amongst the mutants. Received August 14, 2000; Revised and Accepted September 25, 2000. INTRODUCTION In bacteria σ factors are pivotal in establishing regulated and specific initiation of transcription. They bind to the core RNA polymerase and contribute promoter-specific recognition through their DNA binding activities. Additional functions, such as being targets for activator proteins and assisting the DNA strand separation required for RNA synthesis, underline their key roles in gene expression (1). Numerous σ factors are present in bacteria, belonging to two classes. The major class is the σ70 family of proteins; the minor class is represented by σ54 (2,3). The σ54 protein participates in a system of positive control that involves enhancer binding proteins that must hydrolyse a nucleoside triphosphate (NTP) in order to drive the σ54 holoenzyme closed complex to the open complex (4,5). The NTP hydrolysis and enhancer involvement suggest similarities to features of eukaryotic transcription initiation (6). In bacteria, σ54 functions to express genes associated with diverse activities in many different areas of metabolism (reviewed in 7). The domain organisation and functions of σ54 have been explored previously using directed mutagenesis and biochemical characterisation of σ fragments. The domains clearly interact for the full function of σ54 and several distinct activities reside in discrete parts of σ54 (3,8–11). The N-terminal 50 residues (Region I) are required for activator responsiveness and also function to keep the holoenzyme in a transcriptionally silent state prior to activation (5,12–18). Region I directs σ54 to the DNA fork junction that is created when DNA melting starts next to the –12 GC element of the promoter (13,14,19,20). Region I and the C-terminal DNA-binding domain in Region III of σ54 interact (21). Some Region I and II phenotypes are shared and seem to have a common basis in directing fork junction binding and so restricting the conformation of the holoenzyme (22). Protein footprint studies have shown that core polymerase interacts with sequences in Region I and the DNA-binding domain (residues 329–477) suggesting that the interface of σ54 with the core is extensive and specialised (21,23–25). A major core binding determinant is located in Region III between residues 120 and 215 of Klebsiella pneumoniae σ54 (10). Identification of σ54 sequences involved in interactions with promoter DNA, core polymerase and activator provides a basis for understanding the mechanism of transcription initiation. Both directed and random mutagenesis of σ54 have been used. (15–18,26–31). Here we have used random mutagenesis of the full K.pneumoniaerpoN gene, encoding σ54, to identify residues important for functioning of the σ54 holoenzyme in vivo. Subsequently, purified single amino acid substitution mutant proteins were assayed to place limits upon the functional defects in σ54. We identified single amino acids important for functioning of regulatory Region I, for interactions with the core polymerase and for activity of the DNA-binding domain. Mutants defective in transcription initiation but still able to direct closed complex formation were obtained. In addition to its primary core and DNA binding functions, it seems that Region III contributes to activities required for conversion of the closed complex to the open complex. Some of these activities are closely associated with DNA binding preferences and some phenotypes further suggest that the interface of σ54 with core polymerase is extensive. MATERIALS AND METHODS Mutagenesis The low copy number plasmid pMM83 (26) encoding K.pneumoniaerpoN was transformed into the mutator strain Escherichia coli XL1 Red (Stratagene). Cells were grown to saturation in Luria broth twice or four times from a 10% inoculum at 37°C. Total growth time was 24 or 48 h. Randomly mutated pMM83 plasmid DNA was then isolated from each culture and mixed together. Growth screen Mutated DNA was transformed into UNF2792 (26), a K.pneumoniaerpoN mutant strain, selecting with 15 µg/ml chloramphenicol on Luria agar plates at 37°C. Transformants were picked into Luria broth in microtiter plate wells (200 µl) with 15 µg/ml chloramphenicol and grown at 37°C for 24 h. Subsequently, 5 µl of cells were spotted onto a M9 minimal medium plate (32) supplemented with 25 µg/ml histidine and either 0.5 or 1 mg/ml arginine to screen for the aut phenotype. After 24 h incubation at 37°C cells were scored for growth. Immunoblots Transformants displaying the aut phenotype were tested for σ54 protein levels using a polyclonal antibody to E.coli σ54 as described previously (33). Transformants were grown in Luria broth overnight, from a single colony inoculum. Cultures (5 ml) were centrifuged, then resuspended in water to a final volume of 50 µl. An aliquot of 10 µl of concentrated cells was lysed with 10 µl of 2× SDS sample buffer, heated at 95°C and 10 µl of each loaded on denaturing 7.5% SDS–PAGE minigels. Separated proteins were blotted onto PDVF membranes. Anti-σ54 (a gift from A. Ishihama; see also 34) and alkaline phosphatase-conjugated anti-rabbit IgG (Promega) antibodies were used for detection. In vivo promoter activation assays Klebsiella pneumoniae UNF2792 containing pMM83 derivatives was transformed with a K.pneumoniaenifH::lacZ translational fusion reporter plasmid pMB1 (35) or a derivative, pWVC88049 (36), with a higher affinity binding site for the holoenzyme. Cells grown in nutrient broth were used to inoculate 4 ml of NFDM (37) supplemented with histidine (125 µg/ml), 20% glucose as the carbon source and aspartic acid (100 µg/ml) as a poor nitrogen source. They were grown overnight at 30°C. In order to allow derepression of nifA synthesis and activity (22), replica cultures were grown in sealed bijoux. Subsequently, β-galactosidase activity was measured. DNA sequencing Small scale plasmid preparations were made using the Qiagen midi column method and DNA sequences obtained by the ABI prism big dye terminator cycle sequencing ready reaction method (Perkin Elmer) using primers distributed along the coding sequence and upstream of the natural promoter of rpoN. At least two independent sequencing reactions for each strand were performed. Purification of σ54 Wild-type and mutant proteins were prepared as N-terminal His-tagged proteins expressed from pET28b(+) in E.coli B834(DE3) (Novagen). Refolding of the L200P, Q351R and S379F mutants was necessary, since these were found in inclusion bodies following overproduction, and a refolded wild-type protein was included for strict comparison. Refolding was from urea as described (8). Where required, mutants were further chromatographed on a heparin column to remove residual core RNA polymerase. Proteins were finally stored in 10 mM Tris–HCl, pH 8.0, 50% (v/v) glycerol, 0.1 mM EDTA and 1 mM dithiothreitol containing 50 mM NaCl. Protein concentrations were determined using the Bio-Rad Protein Assay kit. Purification of activators Azotobacter vinelandiiNifA and E.coli PspFΔHTH were prepared as N-terminal His-tagged proteins as previously described (23,38) In vitro assays of σ54 function These were conducted as described previously (12,14) using the S.meliloti or K.pneumoniaenifH promoters and E.coli core RNA polymerase (Epicentre Technologies). The heteroduplex DNA sequences used and transcription vectors pMKC28 (for S.melilotinifH) and pNH8 (for K.pneumoniaenifH) have been described (22,39). For reproducibility of data experiments were repeated at least four times. Gel shift assays yielded replicate data within 5% of each experiment. RESULTS Mutagenesis of rpoN Initially we explored the use of a high copy number vector carrying K.pneumoniaerpoN in combination with E.colirpoN mutants to establish a screen for non-functional plasmid borne rpoN alleles. This procedure proved unreliable because of toxicity associated with the high copy number leading to a loss of expression of σ54 through spontaneous formation of stop codons in the early translated sequences (data not shown). The most reliable combination for screening defects in rpoN proved to be the low copy rpoN plasmid pMM83 in K.pneumoniae UNF2792 (26) rather than in E.colirpoN mutants. Utilisation of arginine as a nitrogen source (aut phenotype) in K.pneumoniae (as in E.coli) requires a functional σ54 (40). This phenotype was easily discernible by growth on minimal medium agar plates inoculated from liquid cultures (see Materials and Methods and Fig. 1A). Random mutations were introduced into pMM83 (see Materials and Methods). Amongst 10 000 transformants screened for arginine utilisation, 151 were identified that were defective after retransformation and retesting for arginine utilisation. A final immunoblot screen was used to eliminate from our study mutations that resulted in the production of truncated or unstable σ54, possibly arising through gross losses of structural integrity. Of the 151 candidates, five were found to produce a full-length σ54 protein in immunoblots (Fig. 1A). The remainder did not produce any detectable protein or produced only a truncated protein (data not shown). Only one mutant (later characterised as I23N) had a steady-state level equal to the wild-type, suggesting that for the other mutants reduced levels of σ54 could contribute to the lower in vivo activities measured. However, reductions in σ54 levels do not always correlate with reduced in vivo activities (33). For each of the five defective σ mutants which produced full-length protein the entire rpoN and associated natural promoter region that drives expression of rpoN in pMM83 were sequenced. Each mutant harboured base changes that resulted in a single amino acid substitution in σ54 and DNA sequencing confirmed that the σ54 protein detected by immunoblotting was full-length. The five clones selected exhibited the following changes: ATT→AAT (I23N), CTG→CCG (L200P), CAG→CGG (Q351R), TCC→TTC (S379F) and TCC→TTC (S404F). Mutations at these five positions have not been described previously or have not been extensively characterised (26,27). Locations of the individual amino acid substitutions for each mutant are distributed throughout σ54 (Fig. 1B; see also 41). The mutations map to regions associated with activation, core RNA polymerase binding and DNA binding. This confirms that the strategy was random and hence useful in determination of important novel residues. All substitutions were at residues displaying a high degree of conservation, indicating that important sites had been mutated. Secondary structure predictions strongly indicate that L200 lies in an α-helical secondary structure element. Because of the potentially structurally damaging effects of L200P, a L200A substitution was made for strict comparison. Promoter activation in vivo To extend characterisation of the in vivo phenotype and provide a preliminary quantitative characterisation, the five clones directing the synthesis of full-length σ54 that failed to support the aut phenotype were assessed in a promoter activation assay. Results in Figure 2 show that each mutant gave reduced activation of the K.pneumoniaenifH promoter. They were not rescued by the use of the high affinity σ54 mutant nifH promoter present in pWVC88049 (data not shown; 36). This suggests that several of the mutants did not have simple defects in promoter occupancy, but could be defective in post-promoter binding steps. None of the clones tested elevated the unactivated expression level (assayed under nitrogen replete growth conditions; data not shown) showing that they were all activator-dependent mutants in vivo. A range of activity was identified in the in vivo activation assays. Taking into account the different levels of σ54 (Fig. 1A), it seems that I23N, S379F and S404F have clearly reduced activities, suggesting that different types of mutants were represented. The activity of the I23N mutant was clearly substantially reduced at the K.pneumoniaenifH promoter. This contrasts with the modest defect in the arginine growth test (Fig. 1A) and suggests that certain combinations of activator and promoter can reveal a range of transcription deficiencies for any one mutant σ54. Subsequently, each mutant σ54 was purified as a histidine-tagged protein, together with the wild-type, and characterised by a range of in vitro activity assays. In some cases mutant σ54 proteins were found in inclusion bodies and refolded, together with a refolded wild-type σ54 for strict comparison (see Materials and Methods). Core RNA polymerase binding We used a native gel mobility shift assay to measure binding of the five mutant σ proteins to core polymerase (10). In this assay core polymerase runs as a slow species, the wild-type holoenzyme as a faster single band and the free σ closest to the gel front. Among the mutants, only L200P and S379F behaved differently to the wild-type and displayed a modestly reduced affinity for core polymerase (data not shown). However, the S379F mutant failed to run cleanly into the native gel, suggesting a changed conformation and possibly a reduced availability for binding to core polymerase in vitro. It seems that transcription defects with S379F and L200P might include a contribution from altered core interactions. Activation of transcription in vitro We next determined the in vitro transcription activities of the purified proteins to learn if the purified proteins had activities comparable to those determined in vivo. The ability of the wild-type and mutant holoenzymes to form transcriptionally active open complexes at the K.pneumoniae and S.melilotinifH promoters was measured using supercoiled template DNA. The S.meliloti promoter was used as it provides the basis of many other assays of σ54 function (22,42–44). Two activators, A.vinelandii NifA and E.coli PspFΔHTH, were used to gauge activator-dependent transcription and to relate to the in vivo assays using NifA. Since PspFΔHTH lacks DNA binding activity, its use simplifies DNA gel shift assays (13). In transcription assays dGTP was added as the hydrolysable nucleotide used by the activator, to limit the reaction to formation of an open complex. Subsequently, heparin was added to destroy residual closed complexes and unstable open complexes, together with the four rNTPs to allow initiation and transcript elongation from the preformed stable open complex. Results of these single round transcription assays are shown in Figure 3. Refolded wild-type σ54 and L200A were able to perform activator-dependent transcription at the K.pneumoniae and S.melilotinifH promoters, using either NifA or PspFΔHTH as activator. They produced transcripts to a level comparable to that of wild-type σ54. Mutant Q351R was defective for transcription in all the conditions assayed. Mutants I23N and S379F were unable to transcribe from the K.pneumoniaenifH promoter or from the S.meliloti nifH promoter using NifA as activator. Their activities recovered to 50% of the wild-type activity using S.melilotinifH as template and PspFΔHTH as activator (Fig. 3D). In contrast, mutant S404F was capable of NifA-dependent transcription at the S.meliloti promoter (Fig. 3B). Mutant L200P showed promoter-specific behaviour with some decreased activity in certain transcription assays (compare Fig. 3B and D). Although some mutants showed low levels of in vivo transcription (Fig. 2), the combination of PspFΔHTH and the S.meliloti promoter allowed I23N, Q351R and S379F some increase in transcription levels in vitro (Fig. 3). Overall it appears that the purified mutant proteins have diminished activities in accord with the in vivo activities and protein levels. The results also show that the activities of the mutant σ54 proteins in the transcription assays varied with promoter and activator used, demonstrating that the magnitude of the defects in transcription measured can be specific to the combination of promoter and activator used. DNA binding assays We used a gel shift assay with S.melilotinifH promoter DNA to measure σ and holoenzyme promoter binding. In order to determine the defects associated with each mutation, different templates were used to measure binding of σ and holoenzyme to DNA, as occurs on initial closed complex formation, in the closed complex and in the open complex. Sigma binding to double-stranded promoter DNA. Binding to the S.melilotinifH promoter by the wild-type and mutant σ proteins was measured using double-stranded homoduplex DNA (12) to estimate their abilities to support normal closed complex formation (Fig. 4A). Refolded wild-type, L200A and S404F behaved like wild-type. I23N showed a modestly reduced ability to bind homoduplex DNA, but Q351R and S379F were greatly impaired for binding. With L200P binding appeared to be improved compared to wild-type. These results indicate that mutants I23N, Q351R, S379F and L200P have altered interactions with promoter DNA. Holoenzyme binding to double-stranded promoter DNA. Amongst the mutants, only L200P and S379F behaved differently to the wild-type in their ability to form closed complexes at the double-stranded S.melilotinifH promoter (Fig. 4B). In addition to the standard holoenzyme–DNA conformer, both mutants formed additional conformers with reduced mobilities. It seems that core RNA polymerase can recover some of the lost DNA binding activity seen in the σ–DNA binding assays, particularly for Q351R. In all cases the holoenzyme complexes were disrupted by addition of heparin, which is known to dissociate the σ54 holoenzyme closed complexes (data not shown; see also 10). Only in the presence of activator and a hydrolysable nucleoside triphosphate (dGTP) did heparin-stable complexes form (Fig. 4B). Under these conditions wild-type σ54 holoenzyme produced the most heparin-stable complexes. I23N, L200P, Q351R, S379F and S404F consistently formed fewer complexes stable to the heparin challenge. The pattern of reduced stable complex formation by mutants is similar to the activity patterns observed in in vitro transcription assays. Sigma binding to early melted DNA. In closed promoter complexes a local σ54-dependent DNA opening occurs next to the GC promoter element (19). We judged the ability of the mutants to productively interact with DNA fragments containing this early melted DNA. Binding of wild-type and mutant σ54 proteins to an early melted S.melilotinifH promoter was measured using heteroduplex DNA mismatched at –12/–11, next to the –14/–13 GC element (13,14). Refolded wild-type, L200A, L200P and S404F behaved like wild-type σ54 but I23N, Q351R and S379F were unable to bind (Fig. 5A). This result shows that I23N is distinct amongst the mutants in that it binds double-stranded DNA well (see Fig. 4 above) but fails to bind early melted DNA. In contrast, Q351R and S379F have defects with both template types. They may therefore have defects in a major overall DNA binding determinant. Isomerisation of σ bound to early melted DNA. A stage in open complex formation can be studied without core subunits using DNA pre-opened next to the promoter –12 GC element, so called early melted DNA, σ54 and activator (13,14). σ54 bound to early melted DNA responds to activator in a nucleotide hydrolysis-dependent reaction to form a supershifted DNA complex (ssσ–DNA) in which the DNA has melted and the σ isomerised (13,14). In this assay refolded wild-type and L200A showed similar affinity for early melted DNA and similar efficiency of conversion to the isomerised ssσ–DNA complex using the activator protein PspFΔHTH and hydrolysable nucleotide dGTP (Fig. 5A). I23N, Q351R and S379F failed to bind the early melted DNA and ssσ–DNA complex was not detected after addition of activator and hydrolysable nucleotide (Fig. 5A). Although mutants L200P and S404F showed a similar affinity for early melted DNA to wild-type, the amount of ssσ–DNA complex formed was significantly greater. This same relationship was evident over a range of PspFΔHTH concentrations (data not shown). A second set of assays using early melted K.pneumoniaenifH promoter, the activator protein PspFΔHTH and hydrolysable nucleotide dGTP confirmed and extended the results obtained with S.meliloti early melted DNA (Fig. 5B). The initial σ54–DNA complex with early melted K.pneumoniaenifH promoter DNA was difficult to detect, consistent with the known low binding of σ54 and its holoenzyme to this promoter (42). With σ54 at 10 µM an initial complex was evident (data not shown). However, under activating conditions with 1 µM σ54 the supershifted complex was evident, suggesting that conversion of the initial complex to the isomerised one is rapid. The mutants I23N, Q351R and S379F all failed to give any detectable supershifted complex (Fig. 5B), probably because of poor initial binding (Figs 4 and 5A). The L200P and S404F mutants behave differently on the K.pneumoniae DNA, compared to the S.meliloti early melted DNA (Fig. 5A): in addition to the standard ssσ–DNA complex seen with wild-type σ54 and L200A, L200P and S404F formed an additional conformer with reduced mobility (ssσ*–DNA). The slower moving activator-dependant complex formed with the L200P and S404F mutants and early melted K.pneumoniae nifH DNA could result from either a new σ conformation or from activator recruitment. To explore these possibilities assays were done using 32P-end-labelled PspFΔHTH and unlabelled early melted DNA plus the L200P and S404F mutants. No gel mobility shift of the activator was observed, suggesting that the L200P and S404F mutations result in a new activator-dependent change in complex formation rather than in stable recruitment of activator to the σ–DNA complex (data not shown). Overall, the results with early melted DNA indicate a specific early melted DNA interaction defect in I23N and altered activator responses in L200P and S404F and confirm a strong DNA binding defect in Q351R and S379F. Holoenzyme binding to early melted DNA. To determine whether the mutant proteins would support formation of a closed complex on early melted DNA we next measured binding of holoenzymes to early melted DNA (Fig. 5C). As anticipated from the poor binding of I23N, Q351R and S379F, their holoenzymes bound poorly. Surprisingly, so did the holoenzyme formed with S404F, which alone bound early melted DNA well (Fig. 5A). Holoenzymes formed with L200P, L200A and wild-type σ54 all bound >60% of the DNA. We also measured the stability of the holoenzyme–early melted DNA complexes in a heparin challenge assay (Fig. 5C). Wild-type σ54 holoenzyme binds early melted DNA to give a significant proportion of stable complexes after a 5 min challenge with heparin (14). I23N, L200P, Q351R, S379F and S404F formed holoenzyme complexes with early melted DNA that were significantly less resistant to a heparin challenge (Fig. 5C). In the case of I23N, Q351R and S379F formation of heparin-sensitive complexes correlates with poor binding of the mutant σ to early melted DNA (Fig. 5A). Mutants L200P and S404F are exceptions to this as both of these mutant σ54 proteins bind early melted DNA with similar efficiency to the wild-type. Note that in the heparin challenge a σ–DNA complex forms with the L200P and S404F mutants but not the I23N, Q351R or S379F mutants (Fig. 5C). The presence of a fainter σ–DNA band in the minus heparin lanes for these same two mutants and its virtual absence from the wild-type and L200A lanes further suggests a weaker interaction of S404F and L200P with core RNA polymerase in the assay. Overall, the results with early melted DNA show that the amount of holoenzyme complex and its stability is reduced in mutants I23N, L200P, Q351R, S379F and S404F. The basis for the reductions seems different in that L200P and S404F are not themselves defective in binding early melted DNA whereas the other mutants are. Holoenzyme and σ binding to late melted DNA. Binding of the wild-type and mutant σ proteins (without core) to the –10 to –1 opened DNA followed a pattern very similar to that found on double-stranded DNA (Fig. 4), except for S404F at 2 µM, which showed a 7-fold decrease in affinity for late melted DNA. In open complexes formed with the σ54 holoenzyme, DNA is opened in the region from –10 to –1 (5,12). Mutants of σ54 deregulated for transcription can bind heteroduplex DNA templates opened from –10 to –1 independently of activator and nucleotide hydrolysis because the holoenzyme has isomerised (12,22,33). The holoenzyme formed with wild-type σ54 still requires activator and nucleotide hydrolysis to stably bind such templates (12). To determine whether any of the mutant σ54 proteins were deregulated or changed in their interactions with late melted DNA, the ability of the wild-type and mutant holoenzymes to bind a fully melted S.melilotinifH promoter was measured using heteroduplex DNA with the top strand mismatched from –10 to –1 (12). All mutant holoenzymes bound late melted DNA with similar efficiency as did wild-type holoenzyme and this binding was largely unaffected by addition of activator and GTP (data not shown). However, differences were revealed with a heparin challenge to measure stability (Fig. 6A). As expected, activator and nucleotide were needed for the wild-type holoenzyme to make the greatest number of stable complexes. L200A and S379F behaved similarly. In contrast, I23N and, to a lesser extent, L200P, Q351R and S404F very weakly formed heparin-resistant holoenzymes on late melted DNA in the presence of GTP (the S.melilotinifH mRNA starts 5′-GGG; see 22) but absence of activator. This result suggests that some activator-independent isomerisation of the holoenzyme occurred and was stabilised by GTP. It seems that S404F has a specific defect in binding to late melted DNA which is corrected by core RNA polymerase. Overall, the results suggest that the conformation of the holoenzymes formed with the I23N, L200P and Q351R mutants are different to the wild-type holoenzyme and allow activator-independent interactions with DNA opened for –10 to –1. Activator bypass transcription We next explored whether the mutant σ proteins would support activator-independent transcription from a supercoiled DNA template. With this assay we are able to identify mutants that allow holoenzyme isomerisation without activator. The variation in the standard transcription activation assay employs GTP as the initiating nucleotide and omits activator. Heparin is added after initiation has occurred so as to disrupt unstable complexes, trap stable initiated complexes and limit transcription to a single round. The subsequent addition of the remaining three NTPs thus allows an estimate of the number of stable initiated complexes that have formed from activator-independent open complexes (Fig. 6B). The DNA binding assays with –10 to –1 opened DNA indicated that the I23N, L200P and Q351R mutants were candidates for producing activator-independent transcripts, however, only the I23N holoenzyme produced them. Judged by the lower level of detection, all other mutants and the wild-type holoenzyme formed no more than 50-fold fewer transcripts. The failure of Q351R to support activator-independent transcription may reflect formation of fewer closed complexes or the inability of the holoenzyme to stabilise open DNA unless activated. The failure of L200P may relate to defects in σ interactions with core RNA polymerase. DISCUSSION We have identified five new single amino acid substitutions important for σ54 activity. Our in vitro assays clearly show that several of the mutants alter steps after the initial closed complex forms. Previously, the role of σ54 in the activator-dependent initiation process has been explored using directed mutagenesis of motifs or through random mutagenesis of a restricted set of segments of rpoN (15–18,26–31). In many cases multiple substitutions were required to observe a phenotype and conserved residues proved to be surprisingly tolerant to mutation. Further, inferences that certain residues were important for function were drawn from the apparent non-mutability of particular sequences using a statistical analysis of mutation data (27). In our work altered σ54 proteins defective in supporting growth on arginine and in transcription from the K.pneumoniaenifH promoter were obtained by random mutagenesis of rpoN. The amino acid substitutions leading to defects in σ54 functioning fell in conserved Regions I and III of σ54. Since Region II is largely dispensable for σ54 function and deletions and insertions in Region II lead to modest defects in functionality (45,46), it is likely that there are very few Region II positions where disruptive single amino acid substitutions can occur. Differences in activities we have not directly measured in vitro, such as promoter escape, σ release and rebinding, may also contribute to the lower levels of transcription seen with the mutants in vivo. Some reduction in σ54 holoenzyme levels through reduced σ54 levels are also possible (Fig. 1A), but we note that reduced levels of σ54 are tolerated in in vivo promoter activation assays. Below we discuss the activities of the mutants we obtained in Regions I and III. Region I mutant I23N A number of mutagenesis studies have shown that sequences between residues 15 and 47 in Region I are important for the full function of σ54, but few single substitution mutants have been obtained or characterised and, where they have, these are clustered between residues 25 and 37 (15–17,31,33). Our results show that the I23N mutation results in diminished transcription in response to activator and the acquisition of an activator-independent transcription activity. This result agrees with a patch mutagenesis in which a mutant containing three contiguous alanine substitutions at positions 21–23 showed similar properties (14,33). Mutant I23N is specifically defective in binding to a DNA structure representing the early DNA opening found in closed complexes. This and other evidence leads to the view that the conformational stability of the closed complex is diminished, allowing the I23N holoenzyme to isomerise and engage with melted DNA so as to transcribe without activator and to form stable complexes on heteroduplex templates where DNA is opened from –10 to –1. The defects in activator-dependent transcription can be traced to a reduced activity in the σ isomerisation assay. It seems that I23N contributes both to limiting isomerisation in the absence of activator and to activator responsiveness. Region III mutant L200P The L200P mutation may be structurally damaging since the L200A mutation was indistinguishable from the wild-type in all assays applied to it. Probably, L200P disrupts only a local structure since it retained many of the usual activities of σ54. Assays of core polymerase binding by L200P indicated disruption of a structure that contributes to binding to the core polymerase. We note that L200 is within the 120–215 fragment of σ54 that binds core polymerase with an affinity comparable to that of intact σ54 and that an Fe-BABE form of σ54 derivatised at position 198 is active for cleaving core polymerase (10,24). Many of the defects of the L200P holoenzyme can be rationalised in terms of an altered binding of σ54 to core RNA polymerase. However, L200P also showed altered behaviour in the σ isomerisation assays (evident in formation of a new conformer and increased formation of the isomerised product), suggesting a role for the core binding surfaces of σ54 at steps other than initial formation of the holoenzyme. The defects in open complex formation of the L200P holoenzyme may reflect altered core polymerase to σ contacts important for progression to the open complex where changing interactions between σ54 and core polymerase are envisaged. A mutant L199P of Salmonella typhimiuruim σ54 (31) showed a related set of defects and also led to the conclusion that the sequence in σ54 contributed to a set of interactions after closed complex formation. The results with σ54 are fully consistent with the emerging view that the interface of σ with core polymerase is extensive and functionally specialised (10,23). We note that L200P lies within a domain of σ54 that influences DNA binding (9). Altered properties of L200P evident in DNA binding assays are consistent with this. Region III mutant Q351R The Q351R mutant showed a clear strong and specific defect in overall DNA binding, but core polymerase binding appeared unaffected. Q351 clearly falls within the minimal functional DNA-binding domain of σ54 (residues 329–477; see also 8). Although previously suggested to be unimportant for DNA binding (27), it appears that Q351 makes an important contribution to the activity of the DNA-binding domain of σ54. The properties of the Q351R holoenzyme are readily understood in terms of diminished DNA binding by mutant σ54 restricting levels of closed complex formation. In experiments where the solvent accessibility of σ54 was probed using hydroxyl radical cleavage several protected regions were identified in closed complexes. Strongest DNA-dependent protection from cleavage was at residues 362–386 and 397–432, less strong at residues 305–315 and 325–335 (21). It seems that Q351R could interfere with DNA binding through changing the activity of these or other DNA interacting sequences rather than establishing a direct DNA contact itself. We note that Q350 is implicated in DNA binding by statistical analysis of mutant data (27). Interestingly, Q351R holoenzyme was able to form some heparin-stable complexes independently of activator if the DNA was opened from –10 to –1. This suggests that holoenzyme isomerisation was partly accomplished. Reduced promoter occupancy and/or dissociation of the closed complex on a time scale faster than holoenzyme isomerisation could further explain the transcription defects in Q351R. Region III mutant S379F Two defects were evident in the S379F mutant: one appeared to relate to aggregation, preventing the mutant protein from running into native gels, the other to a marked defect in overall DNA binding. Protein footprints showed that in the closed complex σ54 was protected from hydroxyl radical cleavage at positions 362–386, consistent with a DNA contact being made in the 379 region (21). S379 falls within the helix–turn–helix motif of σ54 where certain mutations are known to reduce promoter occupancy (26,28). Hydrophobic substitutions at position 379 led to diminished transcription in vivo, but the effects of substitution by Phe were not reported. S379F holoenzyme showed reduced promoter DNA binding and did not allow activator-independent transcription. It seems that properties of the S379F mutation are in accord with a role for 379 in setting promoter occupancy rather than restricting polymerase isomerisation in the absence of activator. As discussed for Q351R above, reduced DNA binding by S379F holoenzyme could also contribute to reduced levels of transcripts. Region III mutant S404F The defects in transcription seen with the S404F mutant were clearly due to a step(s) after initial DNA binding since the S404F mutant bound double-stranded DNA and core polymerase well. However, the stability of the S404F holoenzyme on early melted DNA was compromised, but binding of S404F σ54 to the same DNA was not. Moreover, activator-dependent isomerisation of S404F bound to early melted DNA occurred with an efficiency at least as great as that of the wild-type σ54. It seems that S404 is important for an early step in the activator-dependent initiation pathway, which is associated with the interaction the holoenzyme makes with early melted DNA. It seems likely that S404 contributes to holoenzyme isomerisation through being part of the interface between σ54 and core polymerase that is operational when early melted DNA forms. Consistent with this, an altered behaviour of S404F in the σ isomerisation assay was evident, with formation of a new conformer and increased formation of the isomerised product on early melted DNA templates, as also seen with L200P (Fig. 5B). S404 is within a conserved part of the DNA-binding domain, where either Ser or Thr are found adjacent to two invariant Phe residues at positions 402 and 403. Prior directed mutagenesis showed that F402 and F403 were important for holoenzyme function and led to the suggestion that residues 402 and 403 contributed to productive interactions with early melted DNA (47). Results obtained with S404F extend this view by showing that the DNA binding activity to a number of different DNA templates, including early melted DNA, is intact, but that holoenzyme stability is changed on early melted DNA. The changed stability does not lead to activator-independent open complex formation or engagement with melted DNA, suggesting that the inhibitory properties of Region I and R336 that prevent activator bypass activities are largely intact in the S404F mutant (22,47). Although S404F holoenzyme produced a low level of activator-independent, heparin-stable complex on heteroduplex DNA opened from –10 to –1, bypass transcription was not evident (Fig. 6). It seems that S404F might not be able to sufficiently overcome the restriction of isomerisation imposed by its efficient binding to the early melted DNA that forms in closed complexes. Poor binding to early melted DNA, as occurs with I23N, seems necessary to allow sufficient isomerisation for a productive interaction with the transiently melted DNA forming on the supercoiled template (Figs 5 and 6; see also 14). However, S404F alone bound late melted DNA poorly. This suggests that S404F may also be defective in interactions closely associated with late DNA opening, needed for unregulated open complex formation. Protein footprints of σ54 implicate residues between positions 397 and 432 in interactions with core polymerase and with promoter DNA that also involve regulatory Region I (21,23). The close relationship between core-dependent protections seen in the absence of Region I and in the presence of Region I plus promoter DNA suggests that some activities of Region I and the 402–404 patch are linked. The phenotypes of holoenzymes assembled with σ54 mutant at position 402, 403 or 404 are consistent with a defect at an early step in initiation that requires the functionality of Region I (47; this work). This functionality seems to be Region I-directed binding of σ54 to early melted DNA that allows the holoenzyme to assume a heparin-stable complex on such DNA structures (13,14,20). Holoenzyme functionality Overall, our results help identify single amino acids important for σ54 function. In several of the mutants more than one functional defect was evident and some of these were shared by other mutants, arguing that several sequences in σ54 interact for its full function. Probably a large fraction of holoenzyme functioning involves σ54 exchanging information about promoter structure with the core subunits via an extensive set of core–σ54 and σ54–DNA interfaces. The L200P and S404F mutants have altered σ isomerisation properties and some altered holoenzyme properties, suggesting that they could have changes in a communication pathway that links changes in σ to interactions with core polymerase. Current models for activation of the σ54 holoenzyme suggest that the activator engages σ54 bound to early melted DNA as created in closed complexes by the σ54 holoenzyme (13,14,19). A network of interactions involving Region I, core polymerase and DNA contacts is then changed to allow isomerisation of σ54 and holoenzyme and binding to melted DNA. Mutants in σ54 with defects in activator contact could have the phenotype of binding to early melted DNA but failing in isomerisation. No such mutants were found in our screen. Further, none of our mutants were differentially rescued by increasing amounts of activator in the isomerisation assay. Although the mutagenesis was clearly not saturating, it is possible that residues in σ54 that contact the activator have several roles, therefore complicating their identification. Activators of the σ54 holoenzyme lack motifs found in helicases, indicating that activation is via a contact to σ54 rather than through creation of a favourable DNA structure for σ54 binding. ACKNOWLEDGEMENTS This work was supported by funding from the BBSRC to M.B. M.T.G. received a CEC TMR fellowship. We thank M.J. Merrick for pMM83 and UNF2972, D. Studholme, M. Chaney and J. Shumacher for their comments on the manuscript and W. Cannon for oligonucleotides. * To whom correspondence should be addressed. Tel: +44 20 7594 5442; Fax: +44 20 7594 5419; Email: [email protected] Present address: María-Trinidad Gallegos, Departamento de Bioquímica, Biología Molecular y Celular de Plantas, Estación Experimental del Zaidín (CSIC), Profesor Albareda 1, 18008 Granada, Spain View largeDownload slide Figure 1. (A) Growth of K.pneumoniae UNF 2792 on arginine with plasmids containing wild-type σ54 (pMM83) or mutant rpoN genes or with vector (pHSG576). The top panel compares growth of I23N with wild-type and vector. The middle panel compares growth of L200P, L200A, Q351R, S379F and S404F with wild-type and vector. The bottom panel shows the corresponding immunoblot. M, marker (purified σ54 protein). (B) Domain structure of σ54 and its three regions (I–III). The position of each of the random mutants is indicated. X-link, cross linking to promoter DNA; HTH, helix–turn–helix motif; Modulation, domain influencing DNA binding. View largeDownload slide Figure 1. (A) Growth of K.pneumoniae UNF 2792 on arginine with plasmids containing wild-type σ54 (pMM83) or mutant rpoN genes or with vector (pHSG576). The top panel compares growth of I23N with wild-type and vector. The middle panel compares growth of L200P, L200A, Q351R, S379F and S404F with wild-type and vector. The bottom panel shows the corresponding immunoblot. M, marker (purified σ54 protein). (B) Domain structure of σ54 and its three regions (I–III). The position of each of the random mutants is indicated. X-link, cross linking to promoter DNA; HTH, helix–turn–helix motif; Modulation, domain influencing DNA binding. View largeDownload slide Figure 2. Activity of K.pneumoniae UNF 2792 (pMB1) (nifH::lacZ) carrying pMM83 (σ54) or mutant derivatives. β-Galactosidase activities were calculated as a percentage of the wild-type. Means of a minimum of 10 assays are shown, with a relative variation of not more than 0.15 of the net value. View largeDownload slide Figure 2. Activity of K.pneumoniae UNF 2792 (pMB1) (nifH::lacZ) carrying pMM83 (σ54) or mutant derivatives. β-Galactosidase activities were calculated as a percentage of the wild-type. Means of a minimum of 10 assays are shown, with a relative variation of not more than 0.15 of the net value. View largeDownload slide Figure 3.In vitro transcription activity of wild-type and mutant σ54 holoenzymes. Assays were repeated at least eight times to enhance reproducibility and activities were within 15% of the average shown. (A) At the K.pneumoniaenifH promoter in the presence of activator protein NifA at 100 nM. (B) At the S.melilotinifH promoter in the presence of NifA at 100 nM. (C) At the K.pneumoniaenifH promoter in the presence of PspFΔHTH at 100 nM. (D) At the S.melilotinifH promoter in the presence of PspFΔHTH at 100 nM. View largeDownload slide Figure 3.In vitro transcription activity of wild-type and mutant σ54 holoenzymes. Assays were repeated at least eight times to enhance reproducibility and activities were within 15% of the average shown. (A) At the K.pneumoniaenifH promoter in the presence of activator protein NifA at 100 nM. (B) At the S.melilotinifH promoter in the presence of NifA at 100 nM. (C) At the K.pneumoniaenifH promoter in the presence of PspFΔHTH at 100 nM. (D) At the S.melilotinifH promoter in the presence of PspFΔHTH at 100 nM. View largeDownload slide Figure 4. Binding of wild-type and mutant σ54 proteins and their holoenzymes to S.meliloti homoduplex promoter DNA. (A) A constant amount of homoduplex DNA (16 nM) was incubated with increasing amounts of σ54 proteins (0.5, 1 and 2 µM). The percentage of holoenzyme-bound DNA was quantified by phosphorimager analysis and plotted. (B) Gel mobility shift assay showing binding of the wild-type and mutant σ54 holoenzymes (Eσ 100 nM) to homoduplex DNA (16 nM) in the presence of PspFΔHTH (4 µM) and dGTP (4 mM) in the absence of heparin (top). The histograms below indicate the number of initial bound complexes (– Heparin) and those surviving a 5 min heparin challenge (+ Heparin, gel not shown). Black, grey and empty bars are additive and represent the Eσ–DNA, Eσ–DNA* and Eσ–DNA** complexes, respectively. View largeDownload slide Figure 4. Binding of wild-type and mutant σ54 proteins and their holoenzymes to S.meliloti homoduplex promoter DNA. (A) A constant amount of homoduplex DNA (16 nM) was incubated with increasing amounts of σ54 proteins (0.5, 1 and 2 µM). The percentage of holoenzyme-bound DNA was quantified by phosphorimager analysis and plotted. (B) Gel mobility shift assay showing binding of the wild-type and mutant σ54 holoenzymes (Eσ 100 nM) to homoduplex DNA (16 nM) in the presence of PspFΔHTH (4 µM) and dGTP (4 mM) in the absence of heparin (top). The histograms below indicate the number of initial bound complexes (– Heparin) and those surviving a 5 min heparin challenge (+ Heparin, gel not shown). Black, grey and empty bars are additive and represent the Eσ–DNA, Eσ–DNA* and Eσ–DNA** complexes, respectively. View largeDownload slide Figure 5. (A) Binding of wild-type and mutant σ54 proteins to S.meliloti –12/–11 heteroduplex promoter DNA and formation of supershifted complexes. Gel shift assays were conducted with 1 µM protein and 16 nM DNA, in the absence and presence of PspFΔHTH (4 µM) and dGTP (4 mM). The percentage of σ–DNA complex and the supershifted complex (ssσ–DNA) was quantified by phosphorimager analysis. (B) Binding of wild-type and mutant σ54 proteins to K.pneumoniae –12/–11 heteroduplex promoter DNA and formation of supershifted complexes (ssσ–DNA). Assays were conducted as in (A). (C) Holoenzyme mobility shift assays of the wild-type and mutant σ54 holoenzymes (Eσ, 100 nM) with S.meliloti –12/–11 heteroduplex promoter DNA (16 nM), represented by the white bars. Holoenzyme–DNA complexes were challenged with heparin (100 µg/ml) for 5 min prior to loading, represented by the grey bars. View largeDownload slide Figure 5. (A) Binding of wild-type and mutant σ54 proteins to S.meliloti –12/–11 heteroduplex promoter DNA and formation of supershifted complexes. Gel shift assays were conducted with 1 µM protein and 16 nM DNA, in the absence and presence of PspFΔHTH (4 µM) and dGTP (4 mM). The percentage of σ–DNA complex and the supershifted complex (ssσ–DNA) was quantified by phosphorimager analysis. (B) Binding of wild-type and mutant σ54 proteins to K.pneumoniae –12/–11 heteroduplex promoter DNA and formation of supershifted complexes (ssσ–DNA). Assays were conducted as in (A). (C) Holoenzyme mobility shift assays of the wild-type and mutant σ54 holoenzymes (Eσ, 100 nM) with S.meliloti –12/–11 heteroduplex promoter DNA (16 nM), represented by the white bars. Holoenzyme–DNA complexes were challenged with heparin (100 µg/ml) for 5 min prior to loading, represented by the grey bars. View largeDownload slide Figure 6. Bypass activity of σ mutants. (A) Heparin-stable holoenzyme complexes. Binding of the wild-type and mutant σ54 holoenzymes (Eσ, 100 nM) to S.meliloti –10 to –1 heteroduplex DNA (16 nM) in the presence of PspFΔHTH (4 µM) and GTP (4 mM). Heparin (100 µg/ml) was added for 5 min before gel loading. (B) Activator-independent in vitro transcription assay of wild-type and mutant σ54 holoenzymes (100 nM) on the S.melilotinifH promoter (pMKC28, 10 nM). View largeDownload slide Figure 6. Bypass activity of σ mutants. (A) Heparin-stable holoenzyme complexes. Binding of the wild-type and mutant σ54 holoenzymes (Eσ, 100 nM) to S.meliloti –10 to –1 heteroduplex DNA (16 nM) in the presence of PspFΔHTH (4 µM) and GTP (4 mM). Heparin (100 µg/ml) was added for 5 min before gel loading. (B) Activator-independent in vitro transcription assay of wild-type and mutant σ54 holoenzymes (100 nM) on the S.melilotinifH promoter (pMKC28, 10 nM). References 1 Gross,C.A., Chan,C., Dombroski,A., Gruber,T., Sharp,M., Tupy,J. and Young,B. ( 1998) Cold Spring Harbor Symp. Quant. 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Menger, Marcus;Eckstein, Fritz;Porschke, Dietmar

doi: 10.1093/nar/28.22.4428pmid: 11071929

Abstract The dynamics of a hammerhead ribozyme was analyzed by measurements of fluorescence-detected temperature jump relaxation. The ribozyme was substituted at different positions by 2-aminopurine (2-AP) as fluorescence indicator; these substitutions do not inhibit catalysis. The general shape of relaxation curves reported from different positions of the ribozyme is very similar: a fast decrease of fluorescence, mainly due to physical quenching, is followed by a slower increase of fluorescence due to conformational relaxation. In most cases at least three relaxation time constants in the time range from a few microseconds to ~200 ms are required for fitting. Although the relaxation at different positions of the ribozyme is similar in general, suggesting a global type of ribozyme dynamics, a close examination reveals differences, indicating an individual local response. For example, 2-AP in a tetraloop reports mainly the local loop dynamics known from isolated loops, whereas 2-AP located at the core, e.g. at the cleavage site or its vicinity, also reports relatively large amplitudes of slower components of the ribozyme dynamics. A variant with an A→G substitution in domain II, resulting in an inactive form, leads to the appearance of a particularly slow relaxation process (τ ≈200 ms). Addition of Mg2+ ions induces a reduction of amplitudes and in most cases a general increase of time constants. Differences between the hammerhead variants are clearly demonstrated by subtraction of relaxation curves recorded under corresponding conditions. The changes induced in the relaxation response by Mg2+ are very similar to those induced by Ca2+. The relaxation data do not provide any evidence for formation of Mg2+-inner sphere complexes in hammerhead ribozymes, because a Mg2+-specific relaxation effect was not visible. However, a Mg2+-specific effect was found for a dodeca-riboadenylate substituted with 2-AP, showing that the fluorescence of 2-AP is able to indicate inner sphere complexation. Amplitudes and time constants show that the equilibrium constant of inner sphere complexation is 1.2, corresponding to 55% inner sphere state of the Mg2+ complexes; the rate constant 6.6 × 103 s–1 for inner sphere complexation is relatively low and shows the existence of some barrier(s) on the way to inner sphere complexes. Received August 11, 2000; Revised and Accepted September 29, 2000. INTRODUCTION The hammerhead ribozyme is a small catalytic RNA with a core region containing 10 invariant nucleotides and with two substrate binding arms of varying length (1–3). It cleaves the substrate with a certain specificity 3′ to a triplet of the general formula NUH where H can be adenosine, cytidine or uridine but not guanosine. Activity requires divalent metal ions such as Mg2+ or high concentrations of monovalent ions (4). The reaction catalyzed is a phosphoryl transfer where the 2′-OH group of the nucleoside at the cleavage site attacks the phosphorus at the internucleotidic linkage in an in-line mechanism to produce a terminal nucleoside 2′,3′-cyclic phosphate. Several X-ray structures of this ribozyme have been determined, including some structures of intermediate conformations, which probably represent an approach to the transition state (5–7). The general structure is consistent with several solution studies determined mainly by FRET analysis (8–10). Despite this information the structure of the active conformer and details of the reaction mechanism are still elusive. In particular the precise role of the metal ion is yet uncertain. The overall kinetics of the hammerhead ribozymes is dominated by the steps of substrate binding and/or product release (11), which are determined by the well known mechanism of RNA double helix formation and dissociation (12,13). The cleavage rates are Mg2+-dependent and there is good evidence that one of the roles of Mg2+ is to support the formation of the catalytically competent structure. Mg2+-dependent conformational changes have been observed by gel mobility assays and by FRET analysis (9,10). We have previously determined equilibrium constants for Mg2+-induced conformation changes using the fluorescence of 2-aminopurine (2-AP) incorporated at various positions in the ribozyme (14). We report now on the time dependence obtained by fluorescence detected temperature jump relaxation for the same ribozymes as used in the previous study to obtain insight into the internal dynamics of the ribozyme. We have demonstrated that 2-AP can be introduced without loss of catalytic function (14). Substitutions at different positions of the ribozyme facilitate an analysis of the topology of the ribozyme dynamics. The data show that there are both global and local modes of the dynamics, distributed over a broad time range. MATERIALS AND METHODS The hammerhead ribozymes (Fig. 1) and the dodeca-riboadenylate substituted with 2-AP, A(pA)6p(2-AP)(pA)4 [2-AP-oligo(A)], were synthesized by the solid phase method and purified as previously described (14,15). A standard buffer B containing 0.1 M NaClO4, 50 mM cacodylic acid/Tris pH 7.2 was used for most of the measurements. Fluorescence intensities were measured using an SLM 8000 spectrofluorimeter, interfaced to a PC for data collection and averaging. The fluorescence was excited at 320 nm with a bandwidth of 1 nm; the emission was measured at 380 nm with a bandwidth of 16 nm. Before measurements, all solutions were centrifuged within the cuvette in a low-speed desk-top centrifuge for 2 min, in order to clear the solution from dust particles. After thermostating in the cuvette holder for 5 min, the fluorescence intensity was measured for 400 s. There was no indication of a slow process under these conditions. After correction by reference measurements, the data were fitted by least-squares fitting routines using the facilities of the Gesellschaft für wissenschaftliche Datenverarbeitung mbH (Göttingen, Germany). The model used for fitting of the titration data has been described previously (14). Fluorescence detected temperature jump relaxation was measured with an instrument of the type described by Rigler et al. (16) with improvements by C. R. Rabl (unpublished results). A 200 W Xe/Hg arc lamp together with a Schoeffel GM250 monochromator was used as the light source. The fluorescence was excited at 313 nm and collected behind cutoff filters GG385 (2 mm) from Schott (Mainz, Germany). The transients were initially stored on a Biomation 1010 waveform recorder and finally transmitted to the facilities of the Gesellschaft für wissenschaftliche Datenverarbeitung mbH, for exponential fitting by deconvolution procedures either according to Porschke and Jung (17) or Diekmann et al. (18). Stopped flow data were measured with fluorescence detection (excitation/emission wavelengths and cutoff filters as in T-jump experiments) using an instrument developed at the Max Planck Institut für biophysikalische Chemie. The stopped flow construction of this instrument is equivalent to that used in a recently described stopped flow field jump instrument (19). RESULTS Design of hammerhead constructs The hammerhead ribozymes used in the present investigation were cis-cleaving constructs with 49 nt in which cleavage was inhibited by the presence either of 2′-deoxycytidine at position 17 for hammerheads HH1, HH3, HH4 and HH5 or of 2′-deoxy-2′-aminocytidine for HH2 (Fig. 1). The constructs were designed to analyze various parts of the molecule for any characteristic contribution to its dynamics and to metal ion binding. Thus, 2-AP reporter groups were substituted next to the 3′-side of the cleavage site (position 1.1 in HH1, HH2 and HH5), at the catalytic core close to the cleavage site (position 7 in HH3) and at the periphery in the stem III tetraloop (position L3.3 in HH4). Ribozyme HH5 was also modified at position 14, where an adenosine was exchanged by a guanosine, abolishing cleavage activity (22). Broad spectrum of relaxation indicates existence of many conformational substates The general form of the relaxation curves induced by temperature jumps in solutions and reported by the fluorescence of 2-AP is very similar for all the hammerhead ribozyme variants: in a first part of the relaxation curves the fluorescence decreases with time constant(s) below the time resolution of the instrument (τ ≤1 µs); in a second part the fluorescence increases with time constants in the range from a few microseconds up to ~200 ms (Fig. 2). The first part is mainly due to the physical processes of thermal quenching, whereas the second part reflects conformational relaxation of the ribozymes. Exponential fitting of these relaxation curves shows the existence of at least three conformational relaxation processes in most cases. Approximate values of relaxation time constants are compiled in Table 1. An example with at least four conformational relaxation processes required for a satisfactory fit is shown in Figure 3 for HH3. Because of the limited amplitudes and the close spacing of the processes on the time scale, it is virtually impossible to deduce exact values for the time constants. Thus, the existence of a continuous spectrum of time constants cannot be excluded. In spite of this problem in the determination of time constants and the similarity of the general relaxation response, a more detailed comparison of the relaxation curves obtained for the hammerhead variants reveals clear differences (Fig. 2). One extreme case is that of HH4 with 2-AP in the tetraloop, which mainly exhibits a relatively fast relaxation response, which has been observed and characterized previously for isolated hairpin loops (23). The other extreme is HH5, which is not active in catalysis because of the A→G substitution at position 14. In this case the relaxation extends to relatively long times with a rather high amplitude. Thus, it is possible to identify characteristic features of given ribozyme variants. Search for Mg2+-inner sphere complexes Measurements of chemical relaxation can be used for an analysis of metal ion binding with respect to inner sphere complexation. Previous studies (24,25) demonstrated that inner sphere complexation of Mg2+ ions is associated with a characteristic relaxation process in the microsecond time range, which is not observed for Ca2+-ions. Thus, the hammerhead relaxation was monitored over a broad range of Mg2+- and Ca2+-ion concentrations from 0.5 to 100 mM. A sample set of relaxation curves obtained at 5 mM Mg2+ for the various hammerhead species is shown in Figure 4. Addition of Mg2+ ions decreased the relaxation amplitudes and in most cases increased the relaxation time constants. These changes reflect the increased electrostatic shielding resulting from ion binding to the ribozyme. Any additional and specific effect associated with Mg2+-binding, indicating inner sphere binding, has not been detected. However, detection would only be possible if this inner sphere effect would be associated with an amplitude at least as high as the amplitude of the conformational relaxation occurring in the same time range. Because the conformational relaxation extends over the whole time range from a few microseconds to ∼200 ms, it is difficult to get unequivocal evidence for any additional relaxation process in the same time range unless the additional process is associated with a sufficiently high amplitude. Binding of metal ions to the hammerhead ribozyme HH1 was also analyzed by stopped flow experiments with fluorescence detection. Under the conditions of these experiments the fluorescence was affected by a photoreaction and, thus, the stopped flow reaction curves obtained after mixing of ribozyme with Mg2+ had to be corrected by subtraction of the change due to the photoreaction (measured after equilibration of ion binding). Addition of 10 mM Mg2+ to 1.6 µM ribozyme in the standard buffer at 10°C showed the existence of reaction effects with time constants ranging from a few milliseconds over ∼40 s up to ∼20 min (data not shown). These data demonstrate that the spectrum of reaction effects in hammerhead ribozymes extends over a wide time scale. The effects observed upon addition of Ca2+-ions to the hammerhead ribozyme HH1 are very similar to those observed upon addition of Mg2+, both in temperature jump and stopped flow experiments (data not shown). Any clear difference in the data obtained for Mg2+ and for Ca2+ could not be identified. Thus, the Mg2+/Ca2+ comparison, which clearly indicated Mg2+-inner sphere complexation in other cases (cf. below), did not indicate inner sphere complexes in the case of hammerhead ribozymes. Difference relaxation curves illustrate topology of relaxation response The changes of the conformational relaxation observed upon addition of Mg2+ ions provide some information on the nature of the interactions in the hammerhead ribozyme. These changes are more apparent in difference relaxation plots than in the original data. An example is given for HH2 (Fig. 5) using the data for HH1 as a reference: a difference is clearly visible in the absence of Mg2+, but disappears at 5 mM Mg2+. The ribozyme HH2 is distinguished from HH1 by a 2′-NH2-substitution of the nucleotide at the 5′-side of the cleavage site. This substitution leads to a small perturbation of the structure, which is reflected in its relaxation in the absence of Mg2+, but is not visible any more at 5 mM Mg2+. Because of electrostatic repulsion, intramolecular interactions are expected to be more labile in the absence of Mg2+. Addition of 5 mM Mg2+ reduces repulsion, increases the stability of interactions and, thus, masks the perturbation introduced by the 2′-NH2-group. An effect in the opposite direction is found for HH4, where again HH1 is used as a reference. These ribozymes differ in the position of the reporter groups: in HH4 the 2-AP is at the periphery in the stem III tetraloop, whereas in HH1 it is in the core next to the cleavage site. The difference between these two ribozymes is clearly visible in the absence of Mg2+ and increases upon Mg2+ addition (Fig. 6). As mentioned above, the relaxation amplitudes decrease upon Mg2+ addition in all cases. However, the result shown in Figure 6 demonstrates that this decrease is much stronger for HH1 than for HH4. Thus, Mg2+ addition stabilizes the interactions in the core more strongly than those at the periphery. Mg2+-inner sphere complex reported by 2-AP-fluorescence in a substituted dodeca-riboadenylate The relaxation effects obtained for hammerhead ribozymes in the presence of Mg2+ ions and the absence of evidence for formation of inner sphere complexes raises the question whether the fluorescence of 2-AP can be used to detect inner sphere complexes. Because oligoriboadenylic acids are known to form Mg2+-inner sphere complexes (24,25), an oligoriboadenylic acid with 2-AP-substitution was analyzed by the same approach used for the hammerhead ribozymes. Fluorescence detected temperature jump relaxation using 2-AP-oligo(A) revealed a clear relaxation effect with a time constant of ~100 µs in the presence of mM-concentrations of Mg2+ (Fig. 7), which was not observed in the presence of Ca2+ under the same conditions (data not shown). This result demonstrates formation of Mg2+-inner sphere complexes and shows that the fluorescence of 2-AP can be used to detect these complexes. Measurements of the relaxation effect at different Mg2+ concentrations show a dependence of time constants and amplitudes (Fig. 8), which is consistent with the following mechanism Mg2+ + oligo(A)OSC 1 OSCISC 2 In a first reaction step Mg2+ combines with oligo(A) to an outer sphere complex OSC, which is converted in a second, relatively slow reaction to an inner sphere complex ISC. Because the first reaction step is known to proceed without activation barriers as a diffusion controlled reaction, we use k12 = 1 × 1010 M–1 s–1. The overall binding constant K = K1(1 + K2) = (k12/k21)/[1 + (k23/k32)] 3 was determined by fluorescence titrations. The values obtained for Mg2+ and Ca2+ (350 and 320 M–1, respectively) are identical within the limits of accuracy (±100 M–1). Furthermore, the relative change of the fluorescence observed in titration experiments Δ [≡ (fluorescence intensity in the limit of saturation with Me2+)/(fluorescence intensity before addition of Me2+)] is related to the individual quantum yields by Δ = [1/(K2 +1)]α + [K2/(K2 +1)]β 4 where α and β are the quantum yields of OSC and ISC relative to that of the free oligonucleotide, respectively. Again the Δ-values obtained for Mg2+ and Ca2+ (1.30 and 1.26, respectively) are identical within the limits of accuracy (±0.02). Fitting of the data in Figure 8 provided the parameters k23 = 6.6 × 103 s–1 and k32 = 5.5 × 103 s–1. Thus, the inner sphere binding constant K2 = k23/k32 is 1.2 and the outer sphere binding constant K1 is 160 M–1. Due to mutual coupling of the enthalpy changes and the fluorescence quantum yields, these parameters cannot be determined as individual independent values. The combination of these parameters, which has been used to describe the experimental data in Figure 8, is presented in the legend to this figure. DISCUSSION The fluorescence of 2-AP provides a unique approach to the analysis of reaction states and their dynamics in nucleic acids at specific sites. 2-AP may be introduced at various positions within nucleic acid sequences without any serious perturbation of the structure and its fluorescence can be used to report on any process occurring in the environment. It has been shown previously that 2-AP can be introduced into hammerhead ribozymes without loss of catalytic function (14). In the present investigation 2-AP was used to analyze the dynamics and topology of hammerhead ribozymes. Previously the fluorescence of 2-AP has been demonstrated (23) to indicate changes of base stacking interactions in its environment in an oligoriboadenylate with a time constant of <1 µs. Thus, the relaxation effects described here with time constants in the range from a few microseconds to 200 ms must be attributed to internal conformational relaxation of the hammerhead ribozyme. Most of our constructs show at least three relaxation processes which according to theory (26,27) demonstrate the existence of at least four conformational states of the ribozyme. In some cases even more relaxation effects have been observed. Because the relaxation effects are closely spaced on the time scale, it has not been possible to obtain satisfactory evidence for the existence of discrete processes and, thus, a continuous spectrum of relaxation effects reflecting a corresponding spectrum of conformational states cannot be excluded. Although a structural assignment of individual relaxation effects is not possible, the experimental data clearly demonstrate the existence of local modes of conformational relaxation. Hammerhead ribozymes are relatively small RNA molecules and, thus, a general global mode of relaxation may have been expected. However, the experiments show particularly large amplitudes of slow relaxation processes in the catalytic core, whereas fast processes are preferentially reflected from the loop region at the periphery. This investigation complements the earlier study (14) where fluorescence changes of the same 2-AP-containing hammerhead ribozymes were monitored as a function of Mg2+ concentration. The data indicated the presence of multiple binding sites with varying degrees of affinity towards this metal ion. The present results indicate at least three conformational changes at a constant Mg2+ concentration on the pre-steady state time scale. Thus a picture emerges of a rather flexible molecule which undergoes numerous conformational changes to reach its steady state structure. At present it is not possible to assign these changes to a particular structural change. The ribozyme constructs described here are not cleaved and thus additional changes are expected for the cleavage reaction to occur. This is evident from the X-ray structures in the ground state which all require at least one additional local conformational change to reach the transition state to be compatible with an in-line cleavage step (5–7). RNA folding has been the subject of considerable interest (28). In particular, the folding of the domains of the Tetrahymena ribozyme, which is one of the large ribozymes, has also been studied by pre-steady state kinetics. One study employed time-resolved hydroxyl radical footprinting to follow the folding of the entire molecule (29). Another looked at the folding of the P4–P6 subdomains by stopped flow changes of covalently attached pyrene (30). There is a hierarchy of folding events which range from rates of 2 to 0.02 s–1. Thus, the rates in this system are considerably slower than those detected by temperature jump relaxation for the much smaller hammerhead ribozyme. However, except for the fastest folding of the P5a–P5c domain the processes monitored for the Tetrahymena ribozyme are associated with the formation of domain–domain interactions. The resolution of the methods used for the Tetrahymena ribozyme would not have detected the fast internal changes found for the hammerhead ribozyme. In addition to the time range and topology of conformational relaxation, temperature measurements have been used in the present investigation to analyze the type of RNA–Mg2+ interactions. On the basis of phosphorothioate interference studies it is generally accepted that one Mg2+ should be coordinated to the pro-Rp phosphate oxygen at the cleavage site (1,31). Although this has not been shown by X-ray analysis of the ground state structures, a Mg2+ in the vicinity of this position could be identified in the X-ray structure approaching the transition state (6). Using the Rp-diastereomer of a phosphorothioate group at the cleavage site, an inner sphere complex with Hg2+ has been observed by UV–Vis spectroscopy (32). There are a number of other metal ion binding sites detected by uranyl-induced photocleavage (10), NMR spectroscopy (33), X-ray structural analysis (5) and phosphorothioate interference (22,34–36). Molecular dynamics simulations have provided evidence for two Mg2+ ions near the active site (37). A site at G5 previously identified as inhibitory in the presence of Tb(III), by displacing an essential Mg2+, has been further characterized by luminescence spectroscopy (38). These experiments indicate that Tb(III) at this position makes two to three inner sphere contacts with the RNA. Thus, there is evidence for the formation of inner sphere complexation with metal ions other than Mg2+. It would of course be desirable to demonstrate such type of complexation also with this metal ion. Fluorescence detected temperature jump relaxation measurements are suitable to identify the presence of Mg2+-inner sphere complexation on the basis of the characteristic rate constant for inner sphere substitution (25,39,40). This approach has been successful in the analysis of a Mg2+ binding site in the anticodon loop (41) of tRNAPhe and of Mg2+-inner sphere complexes with oligoriboadenylates (24,25). The example described here for the interaction of 2-AP-oligo(A) with Mg2+ as a model system demonstrates that temperature jump relaxation detected by 2-AP fluorescence can distinguish between inner and outer sphere complexation. However, the relaxation spectra of the hammerhead ribozymes were too broad to identify the presence of Mg2+-inner sphere complexation. This does not exclude the existence of such complexes, because their characteristic signal may be hidden in the broad range of conformational relaxation processes. In summary, application of the fluorescence temperature jump technique to hammerhead ribozymes containing 2-AP demonstrates the existence of many different conformational states. The time range of relaxation processes extends from a few microseconds up to 0.2 s. Evidence for a further extension of this time range up to 20 min is provided by stopped flow experiments. Thus, the time scale of conformation changes in hammerhead ribozymes is more extensive than expected. The present investigation also provides clear evidence for the existence of local relaxation modes. This approach should also be useful for the future assignment of relaxation effects to given structure changes. ACKNOWLEDGEMENTS We thank U. Kutzke for her expert preparation of RNA samples. The facilities of the Gesellschaft für wissenschaftliche Datenverarbeitung mbH, Göttingen, were used for data processing. This work was financially supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. * To whom correspondence should be addressed. Tel: +49 551 2011438; Fax: +49 551 2011168; Email: [email protected] View large Download slide View large Download slide Figure 1. (A) Sequence and secondary structure of hammerhead ribozyme variants; numbering according to Hertel et al. (20). (B) Model of the hammerhead ribozyme structure for the sequence used in the present investigation with the positions of substitutions; based on the crystal structure of Pley et al. (21), assuming that the essential parts of the structure remain unchanged. View large Download slide View large Download slide Figure 1. (A) Sequence and secondary structure of hammerhead ribozyme variants; numbering according to Hertel et al. (20). (B) Model of the hammerhead ribozyme structure for the sequence used in the present investigation with the positions of substitutions; based on the crystal structure of Pley et al. (21), assuming that the essential parts of the structure remain unchanged. View largeDownload slide Figure 2. Fluorescence detected relaxation curves (change of fluorescence intensity ΔI as a function of time t) for hammerhead ribozymes HH1–HH5 induced by temperature jumps from 2.0 to 10.6°C in buffer B. The data are shown with a pretrigger recording period of 40 µs, i.e. the time before application of joule heating. The time constant of the initial decrease of the fluorescence is determined by the heating time of 8.25 µs and the detector risetime of 5.4 µs (adjusted in these experiments to a relatively slow detection mode). View largeDownload slide Figure 2. Fluorescence detected relaxation curves (change of fluorescence intensity ΔI as a function of time t) for hammerhead ribozymes HH1–HH5 induced by temperature jumps from 2.0 to 10.6°C in buffer B. The data are shown with a pretrigger recording period of 40 µs, i.e. the time before application of joule heating. The time constant of the initial decrease of the fluorescence is determined by the heating time of 8.25 µs and the detector risetime of 5.4 µs (adjusted in these experiments to a relatively slow detection mode). View largeDownload slide Figure 3. Relaxation curve for HH3 induced by a temperature jump from 2.0 to 10.6°C in buffer B with 5 mM Mg2+, shown in two different time scales (a) and (b). The line marked with circles represents the reference curve measured with a solution of tryptophan under identical experimental conditions; this reference is used for deconvolution (17). The line without noise represents a least squares fit with the time constants 0.56, 5.0, 54, 98 and 8800 µs; the corresponding relative amplitudes are –215.2, 59.7, 18.6, 20.7 and 16.3%. The two lower panels show the residuals for the fast and the slow time scale. The insert shows the autocorrelation function of the residuals. View largeDownload slide Figure 3. Relaxation curve for HH3 induced by a temperature jump from 2.0 to 10.6°C in buffer B with 5 mM Mg2+, shown in two different time scales (a) and (b). The line marked with circles represents the reference curve measured with a solution of tryptophan under identical experimental conditions; this reference is used for deconvolution (17). The line without noise represents a least squares fit with the time constants 0.56, 5.0, 54, 98 and 8800 µs; the corresponding relative amplitudes are –215.2, 59.7, 18.6, 20.7 and 16.3%. The two lower panels show the residuals for the fast and the slow time scale. The insert shows the autocorrelation function of the residuals. View largeDownload slide Figure 4. Fluorescence detected relaxation curves for hammerhead ribozymes HH1–HH5 induced by temperature jumps from 2.0 to 10.6°C in buffer B with 5 mM Mg2+. The data are shown with a pretrigger recording period of 40 µs, i.e. the time before application of joule heating. The time constant of the initial decrease of the fluorescence is determined by the heating time of 7.25 µs and the detector risetime of 5.4 µs (adjusted in these experiments to a relatively slow detection mode). The conditions are identical to those in Figure 2 except for the addition of 5 mM Mg2+. View largeDownload slide Figure 4. Fluorescence detected relaxation curves for hammerhead ribozymes HH1–HH5 induced by temperature jumps from 2.0 to 10.6°C in buffer B with 5 mM Mg2+. The data are shown with a pretrigger recording period of 40 µs, i.e. the time before application of joule heating. The time constant of the initial decrease of the fluorescence is determined by the heating time of 7.25 µs and the detector risetime of 5.4 µs (adjusted in these experiments to a relatively slow detection mode). The conditions are identical to those in Figure 2 except for the addition of 5 mM Mg2+. View largeDownload slide Figure 5. Difference relaxation curves obtained by subtraction of a relaxation curve measured for HH1 from a corresponding curve measured for HH2. (a) Buffer B; (b) buffer B + 5 mM Mg2+. Experimental conditions as described in the legends to Figures 2 and 4. View largeDownload slide Figure 5. Difference relaxation curves obtained by subtraction of a relaxation curve measured for HH1 from a corresponding curve measured for HH2. (a) Buffer B; (b) buffer B + 5 mM Mg2+. Experimental conditions as described in the legends to Figures 2 and 4. View largeDownload slide Figure 6. Difference relaxation curves obtained by subtraction of a relaxation curve measured for HH1 from a corresponding curve measured for HH4. (a) Buffer B; (b) buffer B + 5 mM Mg2+. Experimental conditions as described in the legends to Figures 2 and 4. View largeDownload slide Figure 6. Difference relaxation curves obtained by subtraction of a relaxation curve measured for HH1 from a corresponding curve measured for HH4. (a) Buffer B; (b) buffer B + 5 mM Mg2+. Experimental conditions as described in the legends to Figures 2 and 4. View largeDownload slide Figure 7. Fluorescence detected relaxation curve for 2-AP-oligo(A) in buffer B + 20 mM Mg2+ induced by a temperature jump from 2.0 to 10.6°C. The line marked with circles is the tryptophan reference curve used for deconvolution; least squares fitting provides a value of 95 µs for the chemical relaxation effect. View largeDownload slide Figure 7. Fluorescence detected relaxation curve for 2-AP-oligo(A) in buffer B + 20 mM Mg2+ induced by a temperature jump from 2.0 to 10.6°C. The line marked with circles is the tryptophan reference curve used for deconvolution; least squares fitting provides a value of 95 µs for the chemical relaxation effect. View largeDownload slide Figure 8. Reciprocal relaxation time constants 1/τ (filled circles) and relative fluorescence amplitudes ΔF/F (open circles) obtained by temperature jump measurements for 2-AP-oligo(A) at different Mg2+ concentrations in buffer B at 10.6°C. The dotted and the continuous lines represent a combined fit of these data according the reaction model specified in equations 1 and 2. The parameters are: k12 = 1 × 1010 M–1s–1; k21 = 6.3 × 107 s–1; k23 = 6.6 × 103 s–1; k32 = 5.5 × 103 s–1; α = 0.01; β = 2.38; Δln K1 = 0; Δln K2 = –0.0222. View largeDownload slide Figure 8. Reciprocal relaxation time constants 1/τ (filled circles) and relative fluorescence amplitudes ΔF/F (open circles) obtained by temperature jump measurements for 2-AP-oligo(A) at different Mg2+ concentrations in buffer B at 10.6°C. The dotted and the continuous lines represent a combined fit of these data according the reaction model specified in equations 1 and 2. The parameters are: k12 = 1 × 1010 M–1s–1; k21 = 6.3 × 107 s–1; k23 = 6.6 × 103 s–1; k32 = 5.5 × 103 s–1; α = 0.01; β = 2.38; Δln K1 = 0; Δln K2 = –0.0222. View largeDownload slide Table 1. Relaxation time constants τi of hammerhead ribozymes HH1–HH5 from fluorescence detected temperature jump relaxation measurements The arrows indicate the change of the respective time constants observed with increasing concentration of metal ions: ⇑ = increase of τ (≡ slower relaxation), ⇓ = decrease of τ (≡ faster relaxation); ⇕ = direction not clearly defined. View largeDownload slide Table 1. 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Hoopes, Barbara C.;Bowers, Geoffrey D.;DiVisconte, Matthew J.

doi: 10.1093/nar/28.22.4435pmid: 11071930

Abstract Despite much information as to the structure and function of the general transcription factors, little is known about the regulation of their expression. Transcription of the Saccharomyces cerevisiaeSUA7 (TFIIB) gene results in the formation of two discrete transcripts. It was originally reported that the two transcripts were derived from two promoters separated by ~80 bp. We have found that the two transcripts are instead derived from a common promoter and differ at the 3′-end by ~115 bp. The longer of the two transcripts has an unusually long 3′-untranslated region. We have analyzed the levels of these transcripts under different cell growth conditions and find that the relative amounts of the two transcripts vary. Approximately equal amounts of each transcript are observed during exponential growth, but stresses and growth limiting conditions lead to a decrease in the relative amount of the larger transcript. These results suggest that the expression of the SUA7 gene may be controlled by regulation of 3′-end formation or mRNA stability. One of the general transcription factors, then, may be subject to regulation by a general response of the mRNA processing machinery. Received August 11, 2000; Revised and Accepted September 26, 2000. INTRODUCTION The SUA7 gene of Saccharomyces cerevisiae encodes TFIIB, one of the general transcription factors required for expression of all protein encoding genes (1). This essential transcription factor has two primary roles in the assembly of the initiation complex. First, it is required for the association of the RNA polymerase with the promoter bound transcription factor TFIID by direct contacts with both the TATA binding protein (TBP) and the RNA polymerase (2,3). In addition, clear genetic and biochemical evidence exists that TFIIB is one determinant of start site selection in yeast (4–7). As a critical point in initiation complex assembly, TFIIB has been suggested to be a transcription-limiting step in assembly that can be increased by interaction with activator proteins (8,9). Consistent with this, protein–protein interactions have been detected for TFIIB with a number of activators (10,11) and activation defective mutants of TFIIB have been isolated (12). Addition of TFIIB has been observed to overcome the effect of activator squelching in vitro (13) and in vivo (11). In addition, in vitro mechanistic experiments have suggested that TFIIB addition to the TFIID–TATA complex is a limiting step that can be overcome by the effect of activator proteins (14,15). Although the idea that TFIIB levels play a role in promoter activity and activation is not without controversy (16–18), these other experiments suggest that the concentration of TFIIB in the cell could be important to promoter activity and raise the question of what cellular conditions might affect its intracellular concentration. Despite the potential importance of cellular levels of the general transcription factors, little is known about how their expression might be controlled. The promoters from the human (19), mouse (20), Acanthamoebae (21), Drosophila (22) and yeast (23) TBP genes have been examined. Although these studies identified sequences required for expression in vitro and/or in vivo, they did not address the question of whether cellular conditions affect the expression of the gene. Studies with yeast have found that the levels of both TBP and some of the TATA-associated factors decrease dramatically in post exponential phases of growth (24). Both TFIIB and TBP have been shown to be regulated during some phases of mammalian development (25,26), suggesting that, at least under some conditions, increased amounts of the general transcription factors are required. To address the question of what conditions might normally affect transcription factor expression, we began an analysis of the SUA7 gene in yeast. Analysis of in vivo RNA showed that the SUA7 gene was encoded in two discrete transcripts that differed by about 100 bases in size (5). Start sites for these two transcripts were mapped to two discrete points by primer extension. The more proximal of the two start sites contains a nearly consensus TATA sequence (TATAAAT) at the appropriate distance from the start site. The more distal of the two start sites does not contain a reasonable TATA sequence and is therefore a ‘TATA-less’ promoter. The presence or absence of a TATA box has been shown to impact the interaction of the general transcription factors and RNA polymerase with the promoter and can lead to differences in regulation for the promoter (reviewed in 27,28). In two well-characterized examples (29–31) different types of core promoters contribute to the expression of the genes examined. In both of these cases, evidence exists for differential usage of the two types of core promoters, TATA-containing and TATA-less, under different conditions. To address the question of transcription factor regulation, we examined the effect of different cell growth conditions on the levels of each of the two transcripts for the SUA7 gene. Such an analysis would let us address the possible significance of the two transcripts and of the two promoters proposed to direct their synthesis. MATERIALS AND METHODS Yeast strains and plasmids The haploid strain 23-3a (MATa bar1 leu2 ura3-52 his4 trp1) was obtained from L. Marsh (Albert Einstein College of Medicine, New York, NY). The diploid strain GLM100 was derived from a mating between XLM44-3-27 (L. Marsh) and EG124 (I. Herskowitz, University of California, San Francisco, CA) and has the genotype a/α TRP1/trp1 LEU2/leu2ura3-52/ura3-52 lys2/LYS2 HIS4/his4ade2-100/ADE2 CAN1/can1 (G.L.Matika and B.C.Hoopes, unpublished). The plasmid pBlsc-SUA7 was used for probe synthesis. It was constructed by isolating the ClaI–BamHI restriction endonuclease generated fragment from pDW5462 (5) that contains SUA7 DNA from –789 to +972 relative to the start point of translation. This DNA fragment was ligated into the ClaI and BamHI sites of pBluescript KS+ (Stratagene, Inc.). Standard procedures were used for recombinant DNA techniques (32), media preparation and the growth of yeast strains (33,34). Materials Saturated acid phenol for RNA work (Ambion, Inc.), Nuclease S1, Klenow enzyme, T7 RNA polymerase, ribonucleotides, RQ1 DNaseI (Promega Corporation), deoxyribonucleotides (Boehringer Mannheim), Taq polymerase (Boehringer Mannheim or Sigma Corporation), MSI nylon membrane, formamide, urea, agarose, 40% acrylamide:bis acrylamide (19:1) (Fisher), diethylpyrocarbonate (Aldrich Chemical Corporation, Sigma Corporation), [α-32P]dATP, [γ-32P]ATP and [α-32P]CTP (New England Nuclear/Dupont), SequenaseTM sequencing kit (US Biochemicals) were purchased from the indicated sources. Synthetic DNA Oligonucleotides were purchased from Midland, Inc. or Integrated DNA Technologies and were used without further purification. The sequences of the oligonucleotides were: SUA7-1 (5′-ATCTATGCTCTCCCTAG-3′), Ro-RNAP (5′-CAAAACTAAAGAGTTTGG-3′), 3′ primer (5′-GACGAGCTCGGATCCTGCAGT18VN-3′), 3′ CLMP (5′-GACGAGCTCGGATCCTGCAGT6-3′), SUA7-3′ (5′-ATAACTTACCGGGCGTTG-3′), SUA7-5′ (5′-TATCACCGGATAACAAGAC-3′), RNA-L (5′-ATCTATGCTCTCCCTAG-3′). The SUA7-1 primer is the same primer used for primer extension in (5). The 3′ primer and 3′ CLMP sequences are the same sequences as those described by Russo et al. (35). The 3′ primer is used as a degenerate primer for cDNA synthesis from all mRNA. The 3′ CLMP primer can then be used as an amplification primer for the cDNA synthesized with the 3′ primer. RNA isolation Cells were isolated from 10 ml portions of cultures by centrifugation, rinsed with 1 ml of water, then reisolated by centrifugation and stored at –70°C until use. Total RNA was prepared from frozen cells by the glass bead–phenol method described (34). The final RNA pellet was resuspended in 50 µl of water treated with diethylpyrocarbonate and the concentration determined by UV spectrophotometry. All RNA samples had a 260:280 ratio of greater than 1.7. Northern blots A modification of the procedure described by Rose et al. (33) was used. Either a formaldehyde–agarose gel or a 1× MOPS non-denaturing agarose gel was used for separation of total RNA. Equal amounts of each RNA sample were ethanol precipitated in the presence of sodium acetate. Samples were resuspended by vigorous pipetting in denaturing sample buffer containing formamide and formaldehyde (33) and were heated to 60°C for 10 min before loading on the gel. After electrophoresis, RNA was transferred to a nylon membrane by upward capillary transfer (32) or by using a Turbo Blotter (Schleicher and Schuell) without alkaline denaturation. Nylon membranes were analyzed for ribosomal RNA bands by staining with methylene blue in 0.3 M sodium acetate as described by Herrin and Schmidt (36). Background was reduced by floating the membrane in boiling water until it cooled to room temperature (33), then the residual methylene blue was removed by incubation with shaking in 0.2× SSC, 1% SDS (36). Pre-hybridization and hybridization were carried out as described by Rose et al. (33) except that dextran sulfate was omitted and hybridization was performed at 50°C. The probe used was an anti-sense RNA containing SUA7 sequences from +972 to +618 synthesized from DraI digested pBlsc-SUA7 with T7 RNA polymerase under conditions recommended by the manufacturer. Quantitation of transcript levels was performed on a Storm 820 Phosphorimager (Molecular Dynamics) using the Image Quant software provided by the manufacturer. S1 nuclease analysis The probe used for S1 nuclease analysis was a single stranded DNA containing SUA7 sequence from +24 to –456 generated by extension of the primer SUA7-1 end-labeled with [γ-32P]ATP using single stranded pBlsc-SUA7 DNA as a template and Klenow enzyme as described in (34). After primer extension, the products were digested with BstEII to generate the 3′-end at –456. The labeled probe was purified from the template DNA by separation on a 5% polyacrylamide–7 M urea gel and the labeled product detected by autoradiography. The band was excised and soaked overnight in 0.5 M ammonium acetate, 1 mM EDTA, then the eluted DNA was purified after addition of 10 µg carrier tRNA by phenol extraction and ethanol precipitation and was resuspended in 40 µl TE. Reactions contained 20–30 µg of total RNA and 2 µl labeled probe, and reactions were performed as previously descibed (32) with 1000 U/ml Nuclease S1. After S1 digestion and ethanol precipitation, samples were resuspended in 6 µl water and 4 µl sequencing stop mix was added. Samples were separated by electrophoresis on a 6% polyacrylamide (19:1)–7 M urea sequencing gel. DNA sequence used to determine the size of protected products was generated using a Sequenase kit and conditions recommended by the manufacturer with double stranded pBlsc-SUA7 denatured with alkali as a template and the SUA7-1 oligonucleotide as a primer. RT–PCR reactions RNA used for RT–PCR using two SUA7-specific primers was first treated with DNase I to remove residual chromosomal DNA present in the isolated RNA. The isolated RNA was brought to 1 mM dithiothreitol, 10 mM MgCl2 and 20 mM Tris–HCl pH. 7.5 before the addition of 20 U placental RNase inhibitor and 1 U RQ1 DNase I in a total reaction volume of 50 µl. After incubation for 15 min at 37°C, 25 µl of DNase I stop mix (1.5 M sodium acetate, 50 mM EDTA) was added and the RNA was purified by phenol–chloroform extraction and ethanol precipitation. Total RNA or RNA treated with DNase I (2 µg) was reverse transcribed with AMV reverse transcriptase (10 U/reaction) using buffer and conditions recommended by the manufacturer. The final 20 µl reaction containing cDNA was diluted to 100 µl after heat inactivation of the reverse transcriptase and was used without further purification. PCR reactions (25 µl) contained 2.5–5 µl of the appropriate cDNA, 1 U Taq polymerase, 125 µM of all four deoxynucleoside triphosphates, 1.5 mM MgCl2, and 15 pmol of each primer in the buffer recommended by the manufacturer. Reactions were subjected to a 3 min incubation at 94°C to denature DNA followed by 35 cycles of 94°C (30 s), 50°C (30 s), 72°C (30 s). Analysis of RT–PCR products Products produced as described above were separated by agarose gel electrophoresis (2.5% Nusieve GTG agarose) and the individual products amplified from excised gel slices using Pfu DNA polymerase as per the manufacturer’s instructions (Promega Corporation). The PCR products were purified using Wizard PCR Purification system (Promega Corporation) and ligated into SmaI restricted pBluescript KS+. Ligations were transformed into chemically competent XL10 cells (Stratagene, Inc.) and colonies containing plasmids with inserts were identified by restriction digestion. Insert containing plasmids were sequenced using the Big-Dye Terminator Cycle Sequencing Kit (PE Applied Biosystems) and analyzed on an ABI prism 310 Genetic Analyzer. Analysis of sequences for 3′-end-processing signals was performed using MacVector 6.5 (Eastman Kodak Co.). RESULTS To begin our investigation of the regulation of expression of the TFIIB gene of S.cerevisiae, we analyzed the levels of TFIIB specific transcripts as a function of cellular growth conditions where transcriptional activity is affected. Cultures of haploid yeast were grown to exponential phase, then subjected to one of several conditions. For amino acid starvation, cells were collected by centrifugation, washed with sterile water, then cultured for 48 h in a synthetic media containing nitrogen and glucose but no amino acids or required nucleotides. The yeast strain used was auxotrophic for several amino acids (see Materials and Methods) so this induced an amino acid starvation response (37). Cells were also allowed to grow for 5 days to stationary phase in rich, glucose-containing medium. Cells in stationary phase show a general decrease in protein synthesis and specific changes in gene expression (38). To examine a well-characterized stress response, the cells were subjected to heat shock (39). The temperature of the culture was rapidly raised to 37°C and the culture incubated at this temperature for 2 h. Total RNA was isolated from portions of the cultures and equal quantities of RNA analyzed by northern analysis. As previously reported by Pinto et al. (5), we observed two discrete SUA7-specific transcripts for cells grown under exponential conditions. RNA isolated from the cells is shown in Figure 1A. However, under the stressed conditions, the relative amount of each transcript changes. It is apparent that the different conditions lead to a decrease in the amount of the longer transcript with an increase in the amount of the shorter. Quantitation of the northern analysis is shown in Figure 1B. Although the relative amount of each transcript changes, the absolute levels of total RNA does not change dramatically (data not shown). These results are also observed for diploid cells. RNA isolated from haploid and diploid cells in exponential phase is shown in Figure 2. [The third band marked with an asterisk is only observed under low stringency hybridization conditions and has a mobility consistent with it resulting from hybridization of the probe to 18S ribosomal RNA (data not shown).] We observe only subtle differences for the transcripts isolated from haploid and diploid cells; there may be more of the longer transcript relative to the shorter transcript for the haploid strain (23-3a) compared to the diploid strain (GLM100). As shown in Figure 2, arresting the cell cycle in the G1 phase by the addition of α-factor to the haploid a cells does not result in a similar shift to the shorter transcript. Although in this experiment it appears that the absolute levels of total RNA changes, this effect is not reproducible. To pursue the question of the differential regulation of the two transcripts, we wished to more accurately measure the relative amounts of each transcript under different conditions. Because the two SUA7-specific transcripts are so similar in size, it is difficult to accurately quantify the relative amount of each transcript by northern analysis. We synthesized a strand specific labeled DNA probe containing SUA7 sequences from +24 to –454 relative to the start point of translation to use for S1 nuclease protection experiments. This labeled DNA probe is expected to generate protected products of 68 bp for the short RNA and 148 bp for the long RNA. RNA was isolated from cells for the growth conditions described above and the amount of each 5′-end assessed by S1 nuclease protection (see Materials and Methods). However, as shown in Figure 3A, only one protected product was observed. As demonstrated by the sequencing reactions produced from the same primer used for production of the DNA probe, this product corresponded to that expected for the start site for the short RNA reported by Pinto et al. (5). The position at which we expect to see the product corresponding to hybridization to the long RNA is marked; there is clearly no protected product at that position. A darker exposure of the autoradiogram also showed no evidence of the longer transcript (data not shown). The nuclease protection results suggest that original primer extension assays might not have identified an actual transcriptional start site. We suspected that this might be the case because experiments in our laboratory using total RNA instead of poly A+ RNA yielded large amounts of primer extension product corresponding to the longer RNA, but only small amounts of the product corresponding to the shorter RNA (C.Coffeen and B.C.Hoopes, unpublished). These results were clearly inconsistent with the relative amounts of each transcript observed by northern analysis and with the published work. One possible explanation for our results would be that the shorter primer extension product corresponds to the start site also identified by nuclease protection assays and that the longer primer extension product is the result of DNA synthesis primed from a different RNA template. To test this hypothesis, we performed an RT–PCR analysis of SUA7- specific RNA. Reverse transcriptase was used for cDNA synthesis from RNA samples using the same primer used in this study and others (5) for primer extension assays. The resulting cDNA was then amplified using three different combinations of primers. One reaction contained the primer extension primer and an oligonucleotide with a 5′-end at –421 relative to the translational start site for Sua7p. These primers should only produce a product from genomic DNA that may be contained in the reaction; this region of the SUA7 chromosomal region is not expected to be found in either transcript. As shown in lanes 2, 5 and 8 of Figure 4, products are only observed from reactions containing DNA and not from the cDNA synthesized with the primer extension primer. A second set of primers contained the primer extension primer and an oligonucleotide with a 5′ at –36 relative to the translational start site for Sua7p. This set of primers should generate a 59 bp product from either transcript. As shown in lanes 3, 6 and 9 of Figure 4, this primer set generates a product of the expected size from reactions containing DNA or the synthesized cDNA. The third set of primers contained the primer extension primer and an oligonucleotide with a 5′-end at –69 relative to the translational start site for Sua7p. This set of primers would produce a product of 92 bp from the longer RNA but could not produce a product from the shorter RNA. As shown in lanes 4, 7 and 10 of Figure 4, this primer set only generated a product of appropriate size from DNA; no product was observed for the reaction containing the cDNA for either normal or heat shocked cells. These results support our proposal that the TATA-less start site identified by primer extension (5) does not direct the synthesis of the longer SUA7-specific RNA identified by northern analysis. Since there are clearly two different SUA7-specific transcripts and the results described above suggest that the transcripts have a single 5′-end with a standard TATA-containing promoter, the differences in length must occur elsewhere in the transcript. Since few transcripts in yeast are spliced, the most likely explanation for differences in length for two would be two potential sites for 3′-end formation and polyadenylation. If this were true, we would expect that RT–PCR analysis of the 3′-ends of the transcript would show two discrete products. The relative amounts of these two products should vary for RNA made from cells grown under different conditions. As shown in Figure 5, RT–PCR analysis of the 3′-end of SUA7-specific transcripts confirms this hypothesis. Samples of RNA were used for cDNA synthesis with a primer designed for analysis of 3′-end formation (40). RNA isolated from exponentially grown and amino acid-starved 23-3a cells was used for cDNA synthesis with this 3′ primer. The cDNA was then amplified by PCR with an SUA7-specific primer and a primer containing the arbitrary sequence used as a ‘tag’ for the cDNA. One of the SUA7-specific primers used had a 5′-end at +593 and the other a 5′-end at +1010. As predicted by our hypothesis that the two SUA7-specific transcripts differ at the 3′-end, amplification of cDNA made from RNA isolated from 23-3a cells isolated in exponential phase showed two discrete products for the 3′-end. The two products are ~730 and 880 bases long for the primer with a 5′-end at +593 (Fig. 5B, lane 2) and 275 and 410 bp long for the primer with the 5′-end at +1010 (Fig. 5B, lane 4). For the RNA isolated from the amino acid starved cells, only a single product is obtained whose size corresponds to the smaller product observed for RNA isolated from cells grown to exponential phase (Fig. 5B, lanes 3 and 5). These results confirm our hypothesis that the two SUA7 transcripts differ at the 3′-end and not at the 5′-end as was originally reported. The two polyadenylation sites appear to differ in the degree to which they specify a precise endpoint for the 3′-end. To obtain information that might allow us to understand this difference, we isolated the short and long RT–PCR products and ligated them into a plasmid vector. Individual clones were then sequenced to determine the sequence at which polyadenylation had occurred. These data are summarized in Figure 6. Sequences of 13 individual clones showed the short transcript resulted from 3′-end formation occurring in a very T-rich region encompassing about 28 bases. Sequences of six clones derived from the long RT–PCR product showed 3′-end formation occurring in a cluster of bases occurring about 35 bases downstream of a TA repeat region. Potential 3′-end-processing sequences in this region were identified by searching the 3′-untranslated region (3′-UTR) for the four classes of sequences identified by Graber et al. (41). The results of this analysis are presented in Table 1 and are shown by the boxed sequences in Figure 6. DISCUSSION Our examination of the effect of growth conditions on the transcript levels of the SUA7 gene has led to two main conclusions. First, we have demonstrated that the synthesis of the two SUA7 transcripts is directed by a single promoter and that the transcripts differ in their 3′-UTR by about 115 bases. Secondly, we have shown that the relative amounts of the long and short transcripts vary as a function of cellular growth conditions and stresses. The original observation that the two SUA7 transcripts differed at the 5′-end was prompted by a correspondence between the difference in size of the two transcripts observed on a northern blot and the size of two prominent primer extension products (5). Our primer extension assays showed a much greater amount of the longer than the shorter product despite the fact that northern analysis showed roughly equal amounts of the two transcripts. This suggested to us that one of the two primer extension products might be an experimental artifact. Primer extension assays are prone to several types of artifacts, including premature terminations and extensions of complementary hairpins to produce double stranded products (32,34). Although the latter reaction would result in a longer product, we favor the idea that the longer primer extension product may actually result from hybridization of the primer used for synthesis to a different, more abundant RNA, than that which encodes SUA7. We suggest this because we observe products even larger in size than the prominent longer product, and because the amount of this product observed is not affected by conditions (such as amino acid starvation) that reduce the level of SUA7 transcription (data not shown). Nuclease protection assays support the idea that the more proximal (TATA-containing) start site is used and that the distal start site is not. There are, of course, potential artifacts associated with the use of nuclease protection assays as well (32,34), but we have supported the presence of a single promoter with RT–PCR assays. We see evidence for only the single 5′-end but for two 3′-ends separated by the expected distance. In contrast to the primer extension experiments, the relative amount of the different 3′-ends observed is responsive to cellular growth conditions in a manner predicted by the northern analysis. The existence of two different 3′-ends for SUA7 is interesting in light of the fact that this region contains two closely spaced convergently transcribed genes. SUA7 is transcribed convergently with SRP54 (SRH1) (5), a gene encoding a subunit of the signal recognition particle (42). There are only 275 bp between the translational stop codons for the two genes. As shown in Figure 6, this means that the longer RNA clearly contains a portion of the SRP54 coding sequence. Depending on where the 3′-end for the SRP54 gene is, both transcripts could in fact overlap with the SRP54 transcript. This raises the question of how the effects we observe for the two SUA7 transcripts may affect the expression of SRP54. There are several other pairs of genes for which this question has already been addressed (43,44). These studies have suggested that convergent transcription and antisense RNA production does not affect the regulation of gene pairs. Therefore, although this is an issue that might be pursued, the effects of stress conditions that we see on SUA7 3′-end formation probably do not affect the expression or function of the essential SRP54 gene. Both of the SUA7 transcripts contain the entire coding sequence for TFIIB and the longer transcript contains an additional ∼115 bases of 3′-UTR. Sequences within the 3′-UTR have been shown to affect mRNA stability, mRNA localization and translation (reviewed in 45–47). Any of these characteristics could differ for the two transcripts, affecting the extent to which they contribute to TFIIB protein levels within the cell under different conditions. The longer of the two RNAs is interesting in that its 3′-UTR is quite long compared to the yeast transcripts that have been analyzed so far. Analysis of the size distribution seen for yeast 3′-UTRs shows a narrow distribution peaking at 100 bases (41). It has been proposed that the short length of yeast 3′-UTRs is a consequence of the action of the nonsense-mediated mRNA decay pathway generally involved in mRNA surveillance (48). The question of how (or whether) the longer of the two SUA7 transcripts escapes nonsense-mediated mRNA decay is an interesting one. Although we do not know what functional differences may exist for the two SUA7 transcripts, we do know that the longer of the two transcripts is more sensitive to changes in growth conditions. The amount of the longer transcript decreases when cells are starved for amino acids or other nutrients, when they are grown to stationary phase or when cells are subjected to a 2 h heat shock. We have purposely examined these phenomena under conditions where cells have been subjected to the altered condition for an extended period of time rather than looking at very transient changes. Most of these situations have been shown to result in transcriptional and post-transcriptional effects on gene expression (39,49,50). It is interesting that in the case of the diploid strain under amino acid starvation conditions, the cells are actually only being starved for uracil. However, since limitation for purines is linked to the general amino acid control response (51), starvation for pyrimidines may actually result in a similar response. The changes we have observed in transcript ratios probably reflect a change in polyadenylation site choice. There are other examples of genes for which a change in 3′-end formation occurs so that the proximal signal is used at the expense of the distal one. The CBP1 gene, required for efficient mitochondrial function, has two 3′-ends, one within the coding region of the gene (52). The ratio between the two transcripts changes when cells are grown on glycerol as opposed to glucose, and these changes are thought to be the result of a switch in 3′-end choice (53). This effect is not specific for CBP1, because it is seen for other unrelated genes (54). It is also possible that the two SUA7 transcripts could differ in stability due to the extra sequence present in the longer RNA, which could contain a regulated destabilizing region. Although most destabilizing regions are thought to act constitutively (47), a destabilizing region within the coding region of the c-fos RNA has been proposed to be responsible for rapid degradation of the transcript only upon serum stimulation in mammals (55) and transcripts encoding subunits of the yeast succinate dehydrogenase complex in yeast appear to be regulated at the level of mRNA stability by the presence or absence of glucose in the media (56). The extent to which the two discrete transcripts for the SUA7 gene contribute to Sua7p (TFIIB) protein levels is an important and unanswered question. Experiments that have examined the levels of some of the general transcription factors in yeast have concluded that although TBP protein levels decreased dramatically in cells past exponential phase, the amount of the Toa1p (TFIIA) and Sua7p (TFIIB) proteins remained relatively constant (24). These results are interesting in light of our observation that the relative levels of the two transcripts that encode TFIIB change under different growth conditions. It could be that this change in the relative amounts of the two transcripts is necessary to allow Sua7p protein levels to stay relatively constant under growth and translation-limited conditions. The general transcription factor TFIIB serves a pivotal role in transcription, serving as the protein link between a promoter-bound TFIID and the RNA polymerase holoenzyme. Our studies on the two transcripts that encode TFIIB in the yeast S.cerevisiae have shown that their levels are sensitive to cellular growth conditions. Determining the differences in function for the two SUA7 transcripts will be an essential first step toward an understanding of how cellular stress and growth controls impact the expression of the general transcription factors. ACKNOWLEDGEMENTS We gratefully acknowledge the contribution of Christin Coffeen to early stages of this work and the technical help of Kathleen Baier. This work was supported by grants MCB-9317062 and MCB-9809917 from the National Science Foundation. Automated sequencing was supported by grant DBI-9970204 from the National Science Foundation. * To whom correspondence should be addressed. Tel: +1 315 228 7344; Fax: +1 315 228 7997; Email: [email protected] Present addresses: Geoffrey D. Bowers, Laboratory of Molecular Neuro-Oncology, Department of Neurosurgery, Emory University, Atlanta, GA 30322, USA Matthew J. DiVisconte, Department of Cancer Immunology and AIDS, Dana-Farber Cancer Institute and Department of Pathology, Harvard Medical School, Boston, MA 02115, USA View largeDownload slide Figure 1. Northern analysis of SUA7-specific transcripts as a function of cell growth conditions. Total RNA (15 µg) isolated from cells grown under different conditions was separated by denaturing agarose gel electrophoresis and transferred to a nylon membrane as described in Materials and Methods. (A) Results of phosphorimager visualization of the membrane after hybridization to an anti-sense SUA7 RNA probe. RNA was isolated from cells harvested in exponential phase, after starvation for amino acids, after a heat shock or after growth to stationary phase. (B) Quantitation of relative transcript levels for different conditions. The volume of each transcript was measured using Image Quant software, then the fraction of each transcript making up the total RNA for that condition was calculated. View largeDownload slide Figure 1. Northern analysis of SUA7-specific transcripts as a function of cell growth conditions. Total RNA (15 µg) isolated from cells grown under different conditions was separated by denaturing agarose gel electrophoresis and transferred to a nylon membrane as described in Materials and Methods. (A) Results of phosphorimager visualization of the membrane after hybridization to an anti-sense SUA7 RNA probe. RNA was isolated from cells harvested in exponential phase, after starvation for amino acids, after a heat shock or after growth to stationary phase. (B) Quantitation of relative transcript levels for different conditions. The volume of each transcript was measured using Image Quant software, then the fraction of each transcript making up the total RNA for that condition was calculated. View largeDownload slide Figure 2. Diploid cells show a similar effect of stress. Total RNA (10 µg) isolated from cells grown under different conditions was separated by agarose gel electrophoresis and transferred to a nylon membrane as described in Materials and Methods. (A) Results of autoradiography are shown after hybridization of the membrane to an antisense SUA7 RNA probe. Total RNA was prepared from cells isolated in exponential phase (lanes 1 and 6), after amino acid starvation (lanes 2 and 7), in stationary phase (lanes 3 and 8), after a 2 h heat shock (lanes 4 and 9), and after a 2 h treatment with α-factor (lane 5). (B) Results of methylene blue staining of the membrane before hybridization to demonstrate equal loading of RNA samples. View largeDownload slide Figure 2. Diploid cells show a similar effect of stress. Total RNA (10 µg) isolated from cells grown under different conditions was separated by agarose gel electrophoresis and transferred to a nylon membrane as described in Materials and Methods. (A) Results of autoradiography are shown after hybridization of the membrane to an antisense SUA7 RNA probe. Total RNA was prepared from cells isolated in exponential phase (lanes 1 and 6), after amino acid starvation (lanes 2 and 7), in stationary phase (lanes 3 and 8), after a 2 h heat shock (lanes 4 and 9), and after a 2 h treatment with α-factor (lane 5). (B) Results of methylene blue staining of the membrane before hybridization to demonstrate equal loading of RNA samples. View largeDownload slide Figure 3. S1 protection analysis of SUA7 transcripts. Total RNA was hybridized with a single stranded template strand DNA probe containing SUA7 DNA from +24 to –454 relative to the start of translation. The hybridized region was analyzed by S1 protection as described in Materials and Methods. (A) Autoradiogram showing the protected products after separation on a sequencing gel. The size of the protected products was determined by comparison to the sequencing ladder generated with a primer containing the same 5′-end as the single stranded DNA probe. The positions for the expected short and long RNAs are marked with arrows. RNA was isolated from cells in exponential phase (lane 1), after heat shock (lanes 2 and 5), after amino acid starvation (lanes 3 and 6) and in stationary phase (lane 4). Samples from the amino acid starved cells contained 20 µg of total RNA; the other samples contained 30 µg of total RNA. Lanes 7 and 8 contain labeled probe in the presence and absence of nuclease S1, respectively. (B) A darker exposure of the protected products from the autoradiogram shown in (A). View largeDownload slide Figure 3. S1 protection analysis of SUA7 transcripts. Total RNA was hybridized with a single stranded template strand DNA probe containing SUA7 DNA from +24 to –454 relative to the start of translation. The hybridized region was analyzed by S1 protection as described in Materials and Methods. (A) Autoradiogram showing the protected products after separation on a sequencing gel. The size of the protected products was determined by comparison to the sequencing ladder generated with a primer containing the same 5′-end as the single stranded DNA probe. The positions for the expected short and long RNAs are marked with arrows. RNA was isolated from cells in exponential phase (lane 1), after heat shock (lanes 2 and 5), after amino acid starvation (lanes 3 and 6) and in stationary phase (lane 4). Samples from the amino acid starved cells contained 20 µg of total RNA; the other samples contained 30 µg of total RNA. Lanes 7 and 8 contain labeled probe in the presence and absence of nuclease S1, respectively. (B) A darker exposure of the protected products from the autoradiogram shown in (A). View largeDownload slide Figure 4. RT–PCR analysis of the 5′-end of the SUA7 transcripts. (A) Diagram of the RT–PCR assay used for detection of transcripts differing at the 5′-end. Total RNA isolated from GLM100 cells harvested in exponential phase or after heat shock was treated with DNase I as described in Materials and Methods and used for cDNA synthesis using the SUA7-1 oligonucleotide (SUA7 sequences from +8 to +24) as a primer. This cDNA was then used for PCR using the SUA7-1 primer and either primers a (SUA7-5′), b (RNAL), or c (Ro-f). (B) The products of PCR reactions are shown after separation on a 6% native polyacrylamide gel. Lane 1, a 100 bp DNA ladder; lanes 2–4, cDNA made from exponential cells; lanes 5–7, cDNA made from heat shocked cells; lanes 8–10, plasmid pDW5462. Lanes 2, 5 and 8, primer a; lanes 3, 6 and 9, primer c; lanes 4, 7 and 10, primer b. View largeDownload slide Figure 4. RT–PCR analysis of the 5′-end of the SUA7 transcripts. (A) Diagram of the RT–PCR assay used for detection of transcripts differing at the 5′-end. Total RNA isolated from GLM100 cells harvested in exponential phase or after heat shock was treated with DNase I as described in Materials and Methods and used for cDNA synthesis using the SUA7-1 oligonucleotide (SUA7 sequences from +8 to +24) as a primer. This cDNA was then used for PCR using the SUA7-1 primer and either primers a (SUA7-5′), b (RNAL), or c (Ro-f). (B) The products of PCR reactions are shown after separation on a 6% native polyacrylamide gel. Lane 1, a 100 bp DNA ladder; lanes 2–4, cDNA made from exponential cells; lanes 5–7, cDNA made from heat shocked cells; lanes 8–10, plasmid pDW5462. Lanes 2, 5 and 8, primer a; lanes 3, 6 and 9, primer c; lanes 4, 7 and 10, primer b. View largeDownload slide Figure 5. RT–PCR analysis of the 3′-end of the SUA7 transcripts. (A) Diagram of the RT–PCR assay used for 3′-end determination. Total RNA isolated from exponential phase and amino acid starved 23-3a cells was treated with DNase I and used for cDNA synthesis using the ‘tagged’ 3′ primer shown. SUA7 specific transcripts were then amplified by PCR using the SUA7-specific primers and the tag primer (see Materials and Methods for primer sequences). (B) Products of the RT–PCR reaction were analyzed by electrophoresis on a 2.5% Nusieve agarose gel in TAE and detected by staining with ethidium bromide. Lane 1 is a 100 bp ladder used as a size standard, lane 2 contains products from RNA made from exponential phase cells with the Ro-RNAP primer (5′-end at +594), lane 3 contains products from RNA made from amino acid starved cells with the Ro-RNAP primer, lane 4 contains exponential RNA with the SUA7-3′ primer (5′-end at +1010) and lane 5 contains RNA from starved cells with the SUA7-3′ primer. View largeDownload slide Figure 5. RT–PCR analysis of the 3′-end of the SUA7 transcripts. (A) Diagram of the RT–PCR assay used for 3′-end determination. Total RNA isolated from exponential phase and amino acid starved 23-3a cells was treated with DNase I and used for cDNA synthesis using the ‘tagged’ 3′ primer shown. SUA7 specific transcripts were then amplified by PCR using the SUA7-specific primers and the tag primer (see Materials and Methods for primer sequences). (B) Products of the RT–PCR reaction were analyzed by electrophoresis on a 2.5% Nusieve agarose gel in TAE and detected by staining with ethidium bromide. Lane 1 is a 100 bp ladder used as a size standard, lane 2 contains products from RNA made from exponential phase cells with the Ro-RNAP primer (5′-end at +594), lane 3 contains products from RNA made from amino acid starved cells with the Ro-RNAP primer, lane 4 contains exponential RNA with the SUA7-3′ primer (5′-end at +1010) and lane 5 contains RNA from starved cells with the SUA7-3′ primer. View largeDownload slide Figure 6. Polyadenylation sites for the SUA7 gene. Shown is the DNA sequence of the 3′-UTR of the SUA7 gene as deposited in GenBank as part of the sequencing of Chromsome XVI. The stop codons for the SUA7 and SRP54 genes are indicated in italics. The sites for polyadenylation determined by sequencing of individual plasmids is denoted by the arrows. Potential efficiency, positioning, pre-cleavage and downstream elements are shown as boxed sequences (41). View largeDownload slide Figure 6. Polyadenylation sites for the SUA7 gene. Shown is the DNA sequence of the 3′-UTR of the SUA7 gene as deposited in GenBank as part of the sequencing of Chromsome XVI. The stop codons for the SUA7 and SRP54 genes are indicated in italics. The sites for polyadenylation determined by sequencing of individual plasmids is denoted by the arrows. Potential efficiency, positioning, pre-cleavage and downstream elements are shown as boxed sequences (41). Table 1. Distribution of 3′-end processing signals in the 3′-UTR for SUA7   Efficiency  Positioning  Pre-cleavage  Downstream  Optimal sequence  TATATA  AAAATA  TTTTAT  TTTTCT  Distance found  –80 to –30  –35 to –20  –20 to +5  +5 to +30  RNAS  none  AAAATA  TTTTATTTTTTC  TTTATT  RNAL  TATATA  None  none  none    Efficiency  Positioning  Pre-cleavage  Downstream  Optimal sequence  TATATA  AAAATA  TTTTAT  TTTTCT  Distance found  –80 to –30  –35 to –20  –20 to +5  +5 to +30  RNAS  none  AAAATA  TTTTATTTTTTC  TTTATT  RNAL  TATATA  None  none  none  The optimal sequences are those given by Graber et al. (41). The sequences found for SUA7 are shown in boxes in Figure 6. View Large References 1 Hampsey,M. ( 1998) Molecular genetics of the RNA polymerase II general transcriptional machinery. Microbiol. Mol. Biol. Rev. , 62, 465–503. Google Scholar 2 Buratowski,S. and Zhou,H. ( 1993) Functional domains of transcription factor TFIIB. Proc. Natl Acad. Sci. USA , 90, 5633–5637. 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