Domain swapping in ribonuclease T1 allows the acquisition of double-stranded activity

Dow-Tien Chen and Alan Lin1

Institute of Genetics, National Yang-Ming University, Shih-Pai, Taipei, Taiwan


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
A mutant of ribonuclease T1 (RNase T1), denoted RNase T{alpha}, that is designed to recognize double-stranded ribonucleic acid was created. RNase T{alpha} carries the structure of RNase T1 except for a part of its loop L3 domain, which has been swapped for a corresponding domain from {alpha}-sarcin. The RNase T{alpha} maintains the pleated ß-sheet structure and retains the guanyl-specific ribonuclease activity of the wild-type RNase T1. A steady-state kinetic study on the RNase T{alpha}-catalyzed transesterification of GpU dinucleoside phosphates reveals a slightly reduced Km value of 6.94x10-7 M. When the stranded specificity is examined, RNase T{alpha} catalyzes the hydrolysis of guanine base not only of single-stranded but also, as by design, of double-stranded RNA. The change of stranded specificity suggests the feasibility of using domain swapping to make a substrate-specific ribonuclease. This study suggests that the loop L3 in RNase T1 can be used as a ‘cassette player’ for inserting a functional domain to make ribonuclease of various specificities.

Keywords: {alpha}-sarcin/cassette player/domain swapping/ribonuclease T1/stranded specificity


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Ribonuclease T1 (RNase T1; EC 3.1.27.3) is the best known representative of a large family of homologous microbial ribonucleases (Hill et al., 1983Go; Sevcik et al., 1990Go; Steyaert, 1997Go). It is a small protein that has proven to be a useful model for the study of protein folding and stability (Pace et al., 1988Go; Shirley et al., 1989Go; Pace,1990Go). The protein belongs to the {alpha} + ß class of proteins with several strands of ß-sheet packed against one single, relatively long (4.5 turn) {alpha}-helix (Arni et al., 1988Go; Pace, 1990Go). The ß-sheet is composed of a cohesive structure of four ß-strands which governs the ribonuclease activity and this is commonly found among the ribonuclease family (Hill et al., 1983Go; Arni et al., 1988Go; Sevcik et al., 1990Go; Matinez-Oyanedel et al., 1991; Pace et al., 1991Go; Steyaert, 1997Go). This structural orientation is also characteristically found in fungal ribosome-inactivating proteins such as {alpha}-sarcin (Campos-Olivas et al., 1996Go) and restrictocin (Yang and Moffat, 1996Go), suggesting that the two proteins are related (Kao and Davies, 1995Go; Hwu et al., 2000Go). RNase T1 is a single-stranded ribonuclease with a pronounced specificity for guanine bases (Hill et al., 1983Go; Steyaert et al., 1991Go), whereas, ribosome-inactivating proteins hydrolyze at the purine bases of single- and double-stranded RNA (Endo and Wool, 1982Go). In addition, ribosome-inactivating proteins hydrolyze a wide spectrum of substrates ranging from large molecules such as ribosomes (Endo and Wool, 1982Go) and super-coiled DNA (Ling et al., 1994Go; Cheung et al., 1996Go) to smaller stem-loop oligomer RNAs (Endo et al., 1988Go; Wool et al., 1992Go). It is believed that the variation in substrate specificity of the two ribonucleases is due to differences in their peripheral loops (Kao and Davies, 1995Go, 1999Go; Kao et al., 1998Go; Correll et al., 1999Go; Hwu et al., 2000Go; Garcia-Ortega, et al., 2002Go). Peripheral loop L3 illustrates how a structural difference will result in different functions. The loop L3 of ribonuclease T1 is composed of 16 common amino acid residues and has no assigned function (Pace et al., 1991Go; Steyaert, 1997Go). On the other hand, the loop L3 of {alpha}-sarcin contains a cluster of charged amino acid residues (Campos-Olivas et al., 1996Go; Yang and Moffat, 1996Go) and is known to be involved in the recognition of double-stem RNA structure of the sarcin domain of ribosomal RNA (Correll et al., 1999Go; Hwu et al., 2000Go; Perez-Canadillas et al., 2000Go).

In this study, an engineering mutant of RNase T1, denoted RNase T{alpha}, is put forward. The proposed RNase T{alpha} is suggested from the concept of combining structural elements belonging to different proteins in order to generate proteins with new properties (Bennett et al., 1995Go; Nixon et al., 1997Go; Beguin, 1999Go). Successful examples of such swapping of domains including the triggering of a chemotaxis transduction (Cochran and Kim, 1996Go), the creation of a bifunctional protein that carries ß-lactamase activity in a maltodextrin-binding protein (Betton et al., 1997Go), the making of a new serine protease by sub-domain shuffling (Hopfner et al., 1998Go) and the acquisition of double-stranded DNA binding in cold shock protein (Wang et al., 2000Go). These results have proved the feasibility of the concept. Accordingly, RNase T{alpha}, which is made up of the structure of RNase T1 except that part of its loop L3 domain has been swapped for a corresponding domain from {alpha}-sarcin, was designed to have an add-on double-stranded specificity. The ability of RNase T{alpha} to hydrolyze a single-stranded RNA and double-stranded RNA was examined in this study. The data presented not only suggest that a new ribonuclease can be created by domain swapping but also imply that RNase T1 is a nature enzyme designed to be engineered.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Construction and expression of RNase T{alpha}

The nucleotide sequence of RNase T1 was amplified by polymerase chain reaction (PCR) from genomic DNA extracted from Aspergillus oryzae using the 5' primer 111, 5'-caaacctcgag-3'and the 3' primer 112, 5'gagagagtcccaa-3'. These primers contained XhoI and SalI restriction sites flanking their 5' and 3' termini, respectively. The PCR product was sub-cloned into the polylinker (XhoI and SalI digested) of pBluescript KS (pKS/T1) (Stratagene). After verification of the sequence, the fragment was re-cloned into pGEM (T) at the XhoI or SalI sites to make the pGEM (T)/T1 plasmid.

The chimera protein RNase T{alpha} was designed to carry the structure of RNase T1 with an insertion domain derived from {alpha}-sarcin. Using a PCR-mediated strategy, two primers, the C- and N-primers were used for amplification with pGEM(T)/T1 as the template. The C-primer, 5'-AAGTTTGATTCGAAGAAGCCCAAGGAACCTggtgcgcaccgtctcctcttc-3', carries the 5' end of a sequence that encodes for the loop L3 of {alpha}-sarcin (30 bases in upper case) and the partial coding sequence of 3' end of the ß3 strand of RNase T1 (21 bases in lower case). Similarly, the sequence of the N-primer, 5'-TTCCTTGGGCTTCGAATCAAACTTgtaaacatctccggagctcagaatcg-3, carries the corresponding inserted site with the partial sequence of the loop L3 from {alpha}-sarcin (27 bases in upper case) plus 26 bases (in lower case) that encode for the 3' end of ß2 strand of RNase T1. Both primers contain a BstBI restriction site (underlined) for future self-ligation. The pGEM(T)/T1 plasmid was used as the template for the PCR amplification. The amplification was executed using the following cycling profile: denaturation (95°C for 1 min); annealing (58°C for 1 min); and extension (72°C for 3.5 min). After 25 cycles, PCR amplification yielded a linear DNA fragment of approximately 3.5 kb. The fragment was digested by DpnI restriction enzyme to eliminate endogenous pGEM(T)/T1 plasmid and further digested with BstBI restriction enzyme to generate sticky ends for the PCR fragments. Consequently, the PCR fragment was ligated to make a pGEM/T1-{alpha} plasmid that carried the coding gene for a chimeric protein, denoted RNase T{alpha}. The encoded RNase T{alpha} protein is composed of the structure of RNase T1 with its loop L3 being replaced by 10 amino acids. After the DNA sequence of the chimera gene had been confirmed, the gene was then sub-cloned into a cytosol expression pET28a vector to make a pET28a/T{alpha} plasmid.

The recombinant plasmid pET28a/T{alpha} was propagated in Escherichia coli strain JM109. Cells were grown at 37°C in the presence of kanamycin (50 µg/ml) in LB broth. The expression of recombinant his-tagged RNase T{alpha} was induced by adding IPTG (250 mg/ml) when an A600 value for the culture of 0.3 was reached. The IPTG-induced cells were harvested by centrifugation. His-tagged RNase T{alpha} proteins were purified by the standard procedures of Ni chromatography. Subsequently, the his-tagged RNase T{alpha} was treated with thrombin that cleaved the carboxylic side of arginine residue of his peptide to release a recombinant RNase T{alpha} protein.

Molecular modeling

The tertiary structure of RNase T{alpha} was simulated on the basis of the crystal structure of RNase T1 (Brookhaven PDB entry 1RNT) (Shirley et al., 1989Go; Matinez-Oyanedel et al., 1991; Pace et al., 1991Go). Using energy minimization and different conditions of constraints, the structure of RNase T{alpha} was generated. The secondary structure and the topological folding of RNase T{alpha} were almost the same as those of RNase T1. All calculations were performed on an SGI Origin 2000 computer using the program DISCOVER as implemented in the package InsightII (Molecular Simulation, San Diego, CA).

Determination of ribonuclease activity

The general ribonuclease activity measurements were carried out by RNA-impregnated SDS–PAGE (Hwu et al., 2000Go). Purified recombinant protein was separated electrophoretically using RNA-impregnated SDS–polyacrylamide gel that contained 2.5 mg/ml of large fragments of ribosomal RNA. The presence of RNA fragments in the gel did not interfere with the resolution of protein separation. After electrophoresis, the gel was renatured by incubation in buffer containing 10 mM Tris–HCl, pH 7.4 and 25% 2-propanol (to extract SDS), and this was followed by four washes with incubation buffer without 2-propanol. The gel was then incubated for 30 min at 35°C in the same buffer with vigorous shaking. The ribonuclease activity was detected by staining the gel with 2% Toluidine Blue O in water. A negative staining effect (white in color) against a blue background (the stained RNA) represented the ribonucleolytic action of protein.

The determination of the base specificity was carried out with the same RNA-impregnated SDS-containing polyacrylamide gel system, except that the gel contained monoribonucleotide polymer (poly U, poly A, poly G or poly C) (purchased from Sigma Chemical, St. Louis, MO) instead of RNA fragments.

The kinetic parameters for the transesterification of GpU dinucleoside phosphates (Sigma Chemical) were determined from initial velocities by measuring the increase in absorbance at 280 nm (Steyaert et al., 1991Go). The concentration of GpU dinucleoside phosphate was varied between 10 and 30 µM. Reactions were started by adding different amounts of RNase T1 or RNase T{alpha} and carried out at 35°C in buffer containing 50 mM imidazole, 50 mM NaCl and 2.5 mM EDTA at pH 6.0.

Synthesis of 34-mer oligo RNA

34-mer oligo RNA that mimics the structure of the sarcin domain from 28S rRNA was synthesized using phage T7 polymerase (Promega Biotech) and synthetic DNA oligomers as templates (England et al., 1980Go; Szewczak et al., 1995). The oligomers used were the sequence of T1 promoter and a sequence of 3'-ATTATGCTGAGTGATATCCCTTAGGACGAGTCATGCTCTCCTTGGCGTCCA-5' that carried a complementary T7 promoter (underlined) and the nucleotide of the sarcin domain. They were annealed at 90°C for 3 min followed by cooling on ice. The transcription reaction was carried out at 37°C for 1 h in a solution containing: 40 mM Tris–HCl, pH 8.0, 9 mM MgCl2, 1 mM spermidine, 5 mM dithiothreitol, 1.5 mM each of the four nucleoside triphosphates (Pharmacia P-L Biochemical), 50 nM of the DNA template and 3 units/µl of T7 RNA polymerase. At the end of transcription, the reaction mixture containing the 34-mer oligo RNA was heated at 90°C for 2 min in buffer and renatured by gradual overnight cooling to 4°C to form a secondary structure that imitated the structure of the sarcin domain (the stem and loop structure). The 34-mer RNA was 3' end-labeled with cytidine 3',5'-[{alpha}-32P]bisphosphate according to the published procedure (Huber and Wool, 1986Go).

Digestion of 34-mer RNA and 5S rRNA

The digestion of 3' end-labeled [32P]-34-mer RNA was carried out in a buffer containing 50 mM Tris–HCl, pH 7.6, 50 mM KCl and 5 mM MgCl2 at 30°C for various times and using various amounts of RNase T{alpha}. The cleaved oligo RNA fragments were detected with a phosphoimager. 5S rRNA from E.coli was purchased commercially and end-labeled with 32P on the 3' end accordingly. The digestion of [32P]-5S rRNA was carried out by the same procedure.

Effects of 2'-GMP on the ribonuclease activity

The nucleotide 2'-GMP is known to cognate the active site within the four-stranded pleated ß-sheet (Arni et al., 1988Go). Such cognition causes an inhibition of the ribonuclease activity of RNase T1. Purified recombinant enzyme was incubated with 18S rRNA (obtained from 40S subunits of rabbit reticulocyte ribosomes) in the absence or presence of different amounts of 2'-GMP in buffer containing 50 mM Tris–HCl, pH 7.5, 50 mM KCl, 5 mM MgCl2, 0.1 mM EDTA and 4% glycerol for 15 min at 30°C. The ribonuclease activity of RNase T1 or RNase T{alpha} under the influence of the 2'-GMP was analyzed using 2% agarose gel electrophoresis in TBE with ethidium bromide staining.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Designing, engineering, expression and the structure of RNase T{alpha}

Based on similar structures of RNase T1 and {alpha}-sarcin (Figure 1AGo), the ribonuclease RNase T{alpha} was created by 3D domain swapping (Bennett et al., 1995Go). The RNase T{alpha} carries the structure of RNase T1 except for the partial peripheral loop L3 of the RNase T1 being replaced by a corresponding loop L3 domain of {alpha}-sarcin. This was done by replacing the tetrapeptide containing S–G–G–S (positions 69–72) of RNase T1 with a sequence, KFDSKKPKEN, derived from {alpha}-sarcin (Figure 1BGo). The site of replacement is within a highly conserved region (boxed in Figure 1BGo) where the end residues are either glycine or proline (boxed in Figure 1BGo). This makes the inserted sequence flexible enough to minimize the changes to the overall structure. The essential structure of the pleated ß-sheet is therefore maintained.



View larger version (33K):
[in this window]
[in a new window]
 
Fig. 1. Making RNase T{alpha} by domain swapping. (A) Comparison of the secondary structures of RNase T1 and {alpha}-sarcin: the circle represents the {alpha}-helix structure ({alpha}1); the rectangles are ß-strands (marked ß1, ß2, etc.); and the curved lines are the peripheral loops (L1, L2, L3, L4 and L5) that connect the secondary structures. The loop L3 is a dotted band to illustrate where the swapping takes place in this study. The numbered residues are the active residues that are involved in the ribonuclease activity of the proteins as determined previously (Steyeart, 1997; Kao and Davies, 1999Go; Hwu et al., 2000Go). (B) The amino acid sequence of the loop L3 from {alpha}-sarcin, RNase T1 and RNase T{alpha}. Residues to be swapped are illustrated with bold italic letters. Boxed residues show identical residues in the region of the loop L3 for proteins. Residues are numbered according to their amino acid sequence in the protein. (C) The predicted structure of RNase T{alpha} (yellow backbone ribbon) superimposed on the determined structure of RNase T1 (red). The structure of RNase T1 is depicted from previous data (Shirley et al., 1989Go; Pace et al., 1991Go). The swapped region of RNase T{alpha} is fully displayed using a ball and stick model. The orientation of the loop L5 in the simulated structure is marked with L5. (D) Purity of recombinant RNase T{alpha} (T{alpha}, 1 µg) and RNase T1 (T1, 1 µg) as analyzed by 15% SDS–PAGE. Molecular weight standards (M) are included for comparison.

 
To assess the possible structure change in the new protein, the structure of RNase T{alpha} was simulated using reiterative calculations and animation. Allowances, such as limited substitution or different degrees of restraints, were applied to obtain a model protein with a stable structure. The derived structure of RNase T{alpha} was then compared with the crystal structure of RNase T1 (Matinez-Oyanedel et al., 1991; Pace et al., 1991Go) and the results show almost complete superimposition of RNase T{alpha} structure on the RNase T1 structure except for the swapped domain L3 and loop L5 (Figure 1CGo). The four-strand pleated ß-sheet in RNase T{alpha} remains intact as predicted, but loop L5 is slightly distorted and slanted towards the surface of the molecule (Figure 1CGo). The observed shift in loop L5 seems to be reasonable because it makes space for the extra residues of the new loop L3, which lies next to the loop L5 (Matinez-Oyanedel et al., 1991; Pace et al., 1991Go). Such a structural change may have an effect on the packing density and this may be observable during gel electrophoresis of the protein.

With good results for the structure of RNase T{alpha}, the gene coding for RNase T{alpha} was cloned by genetic manipulation. The amino acid sequence of cloned RNase T{alpha} was confirmed by DNA sequencing. The protein was expressed, purified, released by treatment with thrombin and then analyzed by SDS-PAGE. The recombinant RNase T{alpha} protein appears as a single band (Figure 1DGo) which moves slightly slower (up-shifted) than RNase T1 in an SDS-containing polyacrylamide gel. The observed result is surprising because the difference in electrophoretic mobility does not match the predicted molecular mass of RNase T{alpha} and the difference is greater than the molecular mass of the inserted amino acid residues. Interestingly, such a slower electrophoretic mobility has also been observed in double RNase T1 mutants (Gln25 -> Lys; Glu58 -> Ala) which have a low ribonuclease activity (Shirley et al., 1989Go). Thus, possibilities of changes to the structure that may affect the function of RNase T{alpha} were of deep concern. To allay this concern, structural changes in RNase T{alpha}, particularly changes to the cohesive pleated ß-sheet structure which carries the catalytic center of RNase T1, were analyzed using circular dichroism (CD). The CD measurements showed that RNase T{alpha} has similar CD spectra to RNase T1 (Figure 2Go), indicating that RNase T{alpha} seems to have adopted the same conformation as RNase T1 in solution.



View larger version (22K):
[in this window]
[in a new window]
 
Fig. 2. Spectroscopic characterization of RNase T{alpha}. Circular dichroism spectra in the near-UV region of RNase T{alpha} (•) and RNase T1 ({circ}) at 0.02 µg protein/µl in 10 mM potassium phosphate, pH 7.5, at 25°C.

 
To confirm further that RNase T{alpha} has indeed kept the compact pleated ß-sheet necessary for ribonuclease activity, the response of RNase T{alpha} to the inhibitory effect of 2'-GMP on ribonuclease activity was examined, because 2'-GMP can cognate the pleated ß-sheet of protein and this consequently inhibits the ribonuclease activity (Arni et al., 1988Go; Steyaert, 1997Go). Based on the co-crystal structure of the RNase T1–2'-GMP complex (Arni et al., 1988Go), the guanine of 2'-GMP cooperatively binds to residue of His40 of the ß1 strand and residues of Glu58 (ß2), Arg77 (ß3) and His92 (at the end of the ß4) (Steyaert and Wyns, 1993Go). Mutation at any one of these residues causes a severe loss of ribonuclease activity (Grunert et al., 1991Go; Steyaert, 1997Go), suggesting that maintaining the closed structure integrity of the pleated ß-sheet structure is essential for the ribonuclease. In essence, any mutant of RNase T1 that displays an inhibition by 2'-GMP would indicate the mutant carries the correct ß-sheet structure. Under this criterion, the response of RNase T{alpha} to 2'-GMP was assayed by measuring the hydrolysis of 18S rRNA. The result shows that the hydrolysis of 18S rRNA was completely inhibited at a molar ratio of 2'-GMP to RNase T{alpha} of 1:1 (Figure 3Go). This implies that 2'-GMP can effectively cognate to the pleated ß-sheet of RNase T{alpha}, indicating that the cohesive structure of pleated ß-sheet has been preserved in RNase T{alpha} in the same state as RNase T1.



View larger version (73K):
[in this window]
[in a new window]
 
Fig. 3. Inhibition effect of 2'-GMP on the ribonuclease activity of RNase T{alpha}. Amounts of 4 µM each of RNase T{alpha} and RNase T1 were pre-incubated with different concentrations of 2'-GMP (as indicated at the top of the figure) for 15 min at 30°C, then the process of hydrolysis of 18S rRNA (obtained from rabbit reticulocyte ribosome) was carried out for 15 min at 30°C (as described in Materials and methods). The results of hydrolysis were analyzed on a 2% agarose gel in TBE buffer and visualized with ethidium bromide staining.

 
Base specificity of RNase T{alpha}

The hydrolysis of 18S rRNA described above implies that RNase T{alpha} is an effective ribonuclease of naked RNA. Therefore, the base specificity of RNase T{alpha} was examined next. Using RNA-impregnated SDS–PAGE, it was found that RNase T{alpha}, in addition to hydrolyzing naked RNA (Figure 4AGo), also effectively hydrolyzed polyguanosine (Figure 4BGo), but not polycytidine (Figure 4CGo), polyuridine (data not shown) or polyadenosine (data not shown). These results indicate that RNase T{alpha} has preserved the guanyl specificity of RNase T1.



View larger version (37K):
[in this window]
[in a new window]
 
Fig. 4. Base-specificity of RNase T{alpha}. The ribonuclease activity of RNase T{alpha} (T{alpha}), RNase T1 (T1) and {alpha}-sarcin ({alpha}) against naked RNA (A), poly G (B) and poly A (C) were detected by the procedure of RNA-impregnated SDS–PAGE as described in Materials and methods. Each lane contains 0.5 µg of protein. The ribonucleolytic digestions on poly U and poly C were negative, therefore the results are not given. Negative Toluidine Blue staining was used.

 
When a steady-state kinetic analysis of the RNase T{alpha}-catalyzed transesterification of GpU was carried out, RNase T{alpha} showed a Km value of 6.94x10-7 M with a Kcat of 12.6 s-1which is comparable to the value for RNase T1 (Table IGo). This suggests that RNase T{alpha} is an active ribonuclease.


View this table:
[in this window]
[in a new window]
 
Table I. Specific activities of RNase T1 and RNase T{alpha} against the dinucleoside phosphate GpU
 
Stranded specificity of RNase T{alpha}

As suggested from the initial notion of this study, the question of whether RNase T{alpha} with its swapped domain had enhanced ribonuclease activity towards double-stranded RNA was therefore examined. To assess this experiment, a 3' end-labeled synthetic 34-mer oligo RNA (Figure 5AGo) was used as the substrate for RNase T{alpha}. The 34-mer oligo RNA represents the stem/loop sarcin domain of large subunit ribosomal RNA, which is the best known substrate for {alpha}-sarcin (Endo et al., 1988Go; Correll et al., 1999Go; Perez-Canadillas et al., 2000Go). The result on the hydrolysis of 34-mer oligo RNA by RNase T{alpha} was as predicted. It was found that RNase T{alpha} initially cleaved single-stranded guanine bases at the 5' end (G2 and G3) to produce a 32-mer and a 31-mer, respectively (Figure 5BGo). At increasing concentration, RNase T{alpha} cleaved at single-stranded G21 of the tetra loop in a manner similar to that for {alpha}-sarcin (Endo et al, 1988Go; Cheung et al., 1996Go). Moreover, RNase T{alpha} also hydrolyzed the double-stranded guanine base G23 in addition to G24, a non-Watson–Crick A17:G24 pairing (Correll et al., 1999Go), releasing an 11-mer and a 10-mer, respectively (Figure 5BGo). In a comparative experiment, RNase T1 showed limited ribonuclease activity at the single-stranded guanine bases, acting at G2 and G3 near the 5' end and at G21 within the tetra loop (Figure 5BGo).



View larger version (57K):
[in this window]
[in a new window]
 
Fig. 5. Stranded specificity of RNase T{alpha}: digestion of the 34-mer sarcin RNA. (A) The structure of 34-mer RNA that mimics the sarcin domain of the large subunit 28S rRNA (Correll et al., 1999Go). Nucleotides marked with numbers refer to the position counting from the 5' end of 34-mer RNA. (B) The 3' end-labeled [32P]-34-mer RNA hydrolyzed by {alpha}-sarcin ({alpha}), RNase T{alpha} (T{alpha}) and RNase T1 (T1). Lanes 1–4, digestion with 0.025, 0.05, 0.1 and 0.2 µg of RNase T{alpha}, respectively; lanes 5–7, with 0.05, 0.1 and 0.2 µg of RNase T1, respectively. Lane {alpha} contains 0.2 µg of {alpha}-sarcin. Limited alkaline hydrolysis (Alk. hyd.) of 3' end-labeled [32P]-34-mer RNA was performed for the identification of the cleavage sites. Arrows in (A) indicate the cleavage sites.

 
To validate further the double-stranded activity of RNase T{alpha}, an E.coli 5S rRNA (5S rRNAE.coli) was used as alternative substrate for RNase T{alpha}. Thus, the digestion was carried out with the 3' end-labeled 5S rRNA E.coli. Cutting sites that generated by both RNase T{alpha} and RNase T1 were assessed against the secondary and tertiary structure of 5S RNA (Brunel et al., 1991Go; Ban et al., 2000Go; Mueller et al., 2000Go) (Figure 6A and CGo). Both ribonucleases limit their hydrolytic activity to guanine bases (Figure 6BGo). However, an examination of the stranded specificity showed that the guanine bases of region II that form a stable helix structure (helix 2) (Brunel et al., 1991Go; Mueller et al., 2000Go) were cleaved by RNase T{alpha}, but not by RNase T1 (Figure 6BGo). Similarly, guanine bases in regions III (helix 3) and IV (helix 4) were only susceptible to hydrolysis by RNase T{alpha}. Again, RNase T{alpha} was active at the reactive guanine bases that undergo non-Watson–Crick pairing (G18:U65; U22:G61; U80:G96) (Brunel et al., 1991Go; Mueller et al., 2000Go) (Figure 6A and CGo). The molecular meaning of the interaction between the non-Watson–Crick pairings and RNase T{alpha} remains to be determined, but it is clear that RNA with a highly organized helix structure is susceptible to RNase T{alpha} (Figure 6CGo). Taken together, these results confirm that RNase T{alpha} is capable of hydrolyzing double-stranded RNA. Hence the idea of using the strategy of domain swapping to create an add-on-function to a ribonuclease T1 has been successful.



View larger version (40K):
[in this window]
[in a new window]
 
Fig. 6. Stranded specificity of RNase T{alpha}: digestion of the 5S rRNA. (A) The secondary structure of 5S rRNA of E.coli as depicted from the studies of Brunel et al. (Brunel et al., 1991Go). (B) The hydrolysis of 5S rRNA by RNase T{alpha} and RNase T1 analyzed by SDS–PAGE on 12% polyacrylamide gel containing 6 M urea. The 5S rRNA was from E.coli and end-labeled with 32P on the 3' end. It was digested with different amounts (1x, 0.02 µg; 10x, 0.2 µg) of RNase T{alpha} and RNase T1. Cleavage sites are marked with numbers that refer to the position of guanine bases counting from the 5' end of 5S rRNA. The accessed guanine bases that are double-stranded are marked with a single asterisk. Cleavages occurring at non-Watson–Crick pairings are marked with a double asterisk. Symbol {triangleup} indicates guanine bases (G83 to G86) that are located in the sensitive double-stranded region that are known to be easily attacked by hydroxyl radicals (Hancock and Wagner, 1982Go; Zhong and Kallenback, 1994). (C) The tertiary locations of reactive guanine bases that are hydrolyzed by RNase T{alpha} are marked with their number. The tertiary structure (illustrated by a backbone ribbon) of E.coli 5S rRNA comes from the work of Mueller et al. (Mueller et al., 2000Go). The helix region is dark and the non-helix region the lighter of the two. Cleavages occurring in the helix I region are not shown.

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The rationale for this study is based on the similarity of the tertiary structures found between RNase T1 (Matinez-Oyanedel et al., 1991; Pace et al., 1991Go) and {alpha}-sarcin (Campos-Olivas et al., 1996Go, Yang and Moffat, 1996Go; Perez-Canadillas et al., 2000Go) (Figure 1AGo). The loop L3 of {alpha}-sarcin and that of RNase T1 are oriented in a very similar geometry relatively to the neighboring domains (ß2 and ß3 for RNase T1; ß4 and ß5 for {alpha}-sarcin) and exposes on the surface of the molecule (Matinez-Oyanedel et al., 1991; Pace et al., 1991Go; Campos-Olivas et al., 1996Go, Yang and Moffat, 1996Go). Theoretically, by swapping the loop L3 of {alpha}-sarcin with the corresponding position of RNase T1, a new RNase should keep its structural integrity and consequently maintain ribonuclease activity. Practically, in this study, the engineered RNase T{alpha} did keep a similar structure to RNase T1. The simulated molecular structure found initially that RNase T{alpha} had a comparable structure to RNase T1 and this was further supported by the physical CD profile of both ribonucleases. Moreover, the biochemical data on the 2'-GMP inhibitory effect showed that RNase T{alpha} carries an intact pleated ß-sheet despite having several extra amino acid residues inserted in the middle of the ß-sheet. With this assured structure, the ribonuclease activity of RNase T{alpha} was also found to be well preserved. The engineered RNase T{alpha} not only displays all the enzymatic properties that RNase T1 does, but also has its strand specificity extended from single-stranded to both single- and double-stranded specificity. The new double-stranded property is assumed to derive from the swapped loop L3 which is known to interact with the helix–stem structure of RNA from the sarcin domain of ribosomal RNA (Correll et al., 1999Go; Hwu et al., 2000Go). Hence the preconceived notion on engineering of a new RNase with an add-on functional domain has been completely demonstrated in this study.

The success of engineered RNase T{alpha} is significant in many aspects. Practically, the engineered RNase T{alpha} is a potential tool for footprinting the structure of ribonucleic–protein complexes (RNP). The use of {alpha}-sarcin with its lack of strand preference and purine specificity has helped to elucidate in detail the structures of the 5S rRNP (Huber and Wool, 1986Go), the 7S rRNP (Sands and Bogenhagen, 1991Go) and the SRP (signal recognition particle) (Stub et al., 1991). Ironically, the properties of the engineered RNase T{alpha}, with it lack of strand preference and guanyl specificity, make RNase T{alpha} a better tool than {alpha}-sarcin for footprinting the RNP complex. An experiment using of RNase T{alpha} to dissect an RNP complex is in progress.

In essence, the most significant aspect of this study, besides proving the feasibility of making a new ribonuclease by domain swapping, is the discovery that the part of the loop L3 in the RNase T1 can be exchanged without jeopardizing its ribonuclease activity. In numerous studies of RNase T1, the activities of single, double or triple mutations on RNase T1 have been reported (Grunert et al., 1991Go; Steyaert and Wyns, 1993Go; Loverix et al., 1997Go, 1998Go; Steyaert, 1997Go; Hubner et al., 1999Go); however, such an exchange of a partial domain has never previously been carried out. Our example of an engineered RNase T{alpha} suggests that RNase T1 is an enzyme capable of further engineering. The potential use of this work can be predicted where RNase T1 becomes a ‘cassette player’ and the loop L3 is a site for where new molecular ‘cassettes’ are inserted. This will allow the creation of a series of novel enzymes with new activities. Any functional module could be considered as a cassette to slot into the position of the loop L3 of RNase T1. Then that RNase T1 would be expected to ‘play’ the activity of an inserted module along with the original ribonuclease activity. Ultimately, in the future it should be possible to generate a wide range of specific ribonucleases using this concept.


    Notes
 
1 To whom correspondence should be addressed. e-mail: alin{at}ym.edu.tw Back


    Acknowledgments
 
We thank Professor R.Kirby of Rhodes University, South Africa, for critical reading of the manuscript. This work was supported in part by grants from the National Science Council (NSC89-2320-B010-128).


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Arni,R., Heinemann,U., Tokuoka,R. and Saenger,W. (1988) J. Biol. Chem., 263, 15358–15368.[Abstract/Free Full Text]

Ban,N., Nissen,P., Hansen,J., Moore,P.B. and Steitz,T.A. (2000) Science, 289, 905–920.[Abstract/Free Full Text]

Beguin,P. (1999) Curr. Opin. Biotechnol., 10, 336–340.[CrossRef][ISI][Medline]

Bennett,M.J., Schlunegger,M.P. and Eisenberg,D. (1995) Protein Sci., 4, 2455–2468.[Abstract/Free Full Text]

Betton,J.M., Jacob,J.P., Hofnung,M. and Broome-Smith,J.K. (1997) Nature Biotechnol., 15, 1276–1279.[Medline]

Brunel,C., Romby,P., Westhof, E., Ehresmann,C. and Ehresmann,B. (1991) J. Mol. Biol., 221, 293–308.[CrossRef][ISI][Medline]

Campos-Olivas,R., Bruix,M., Santoro,J., Martinez del Pozo,A., Lacadena,J., Gavilanes,J.G. and Rico,M. (1996) FEBS Lett., 399, 163–165.[CrossRef][ISI][Medline]

Cheung,J-I., Wang,Y-R. and Lin,A. (1996) FEBS Lett., 386, 60–64.[CrossRef][ISI][Medline]

Cochran,A.G. and Kim,P.S. (1996) Science, 271, 1113–1116.[Abstract]

Correll,C.C., Wool,I.G. and Munishkin, A. (1999) J. Mol. Biol., 292, 275–287.[CrossRef][ISI][Medline]

Endo,Y. and Wool,I.G. (1982) J. Biol. Chem., 257, 9054–9060.[Abstract/Free Full Text]

Endo,Y., Chan,Y-L., Lin,A., Tsurugi,K. and Wool,I.G. (1988) J. Biol. Chem., 263, 7917–7920.[Abstract/Free Full Text]

England,T.E., Bruce,A.G. and Uhlenbeck,O.C. (1980) Methods Enzymol., 65, 65–74.[Medline]

Garcia-Ortega,L., Masip,M., Mancheno,J.M., Onaderra,M., Lizarbe,MA., Garcia-Mayoral,M.F., Bruix,M., Martinez del Pozo,A. and Gavilanes,J.G. (2002) J. Biol. Chem., 277, 18632–18639.[Abstract/Free Full Text]

Grunert,H.P., Zouni,A., Beineke,M., Quaas,R., Georgalis,Y., Saenger,W. and Hahn,U. (1991) Eur. J. Biochem., 197, 203–207.[Abstract]

Hancock,J. and Wagner,R. (1982) Nucleic Acids Res., 10, 1257–1269.[Abstract]

Hill,C., Dodson,G., Heinemann,W., Saenger,W., Mitsui,Y., Nakamura,K., Borisov,S., Tischenko,G., Polyakov,K. and Pavlovsky,S. (1983) Trends Biochem. Sci., 8, 364–369.[CrossRef][ISI]

Hopfner,K.P., Kopetzki,E., Kresse,G-B., Bode,W., Huber,R. and Engh,R.A. (1998) Proc. Natl Acad. Sci. USA, 95, 9813–9818.[Abstract/Free Full Text]

Huber,P.W. and Wool,I.G. (1986) Proc. Natl Acad. Sci. USA, 83, 1593–1597.[Abstract]

Hubner,B. Haensler,M. and Hahn,U. (1999) Biochemistry, 38, 1371–1376.[CrossRef][ISI][Medline]

Hwu,L., Huang, K-C., Chen,D-T. and Lin,A. (2000) J. Biomed. Sci., 7, 420–428.[CrossRef][ISI][Medline]

Kao,R. and Davies,J. (1995) Biochem. Cell Biol., 73, 1151–1159.[ISI][Medline]

Kao,R. and Davies,J. (1999) J. Biol. Chem., 274, 12576–12582.[Abstract/Free Full Text]

Kao,R., Shea,J.E,, Davies,J. and Holden,D.W. (1998) Mol. Microbiol., 29, 1019–1027.[CrossRef][ISI][Medline]

Ling,J., Liu,W.Y. and Wang,T.P. (1994) FEBS Lett., 345, 143–146.[CrossRef][ISI][Medline]

Loverix,S., Doumen,J. and Steyaert,J. (1997) J. Biol. Chem., 272, 9635–9639.[Abstract/Free Full Text]

Loverix,S., Winquist,A.W., Stromberg,R. and Steyaert,J. (1998) Nature Struct. Biol., 5, 365–368.[ISI][Medline]

Martinez-Oyanedel,J., Choe,H.W., Heinemann,U. and Saenger,W. (1991) J. Mol. Biol., 222, 335–352.[ISI][Medline]

Mueller,F., Sommer,I., Baranov,P. Matadeen,R., Stoldt,M., Wohnert, J., Gorlach,M., van Heel,M. and Brimacombe,R. (2000) J. Mol. Biol., 298, 35–59.[CrossRef][ISI][Medline]

Nixon,A.E., Warren,M.S. and Benkovic,S.J. (1997) Proc. Natl Acad. Sci. USA, 94, 1069–1073.[Abstract/Free Full Text]

Pace,C.N. (1990) Trends Biochem. Sci., 15, 14–17.[CrossRef][ISI][Medline]

Pace,C.N., Grimsley,G.R., Thomson,J.A. and Barnett,B.J. (1988) J. Biol. Chem., 263, 11820–11825.[Abstract/Free Full Text]

Pace,C.N. Heinemann,U., Hahn,U. and Saenger,W. (1991) Angew. Chem., Int. Ed. Engl., 30, 343–360.[ISI]

Perez-Canadillas,J.M., Santoro,J., Campos-Olivas,R., Lacadena,J., Martinez del Pozo,A., Gavilanes,J.G, Rico,M. and Bruix,M. (2000) J. Mol. Biol., 299, 1061–1073.[CrossRef][ISI][Medline]

Sands,MS. and Bogenhagen,D.F. (1991) Nucleic Acids Res., 19, 1797–1803.[Abstract]

Sevcik,J., Sanishvili,R.G., Pavlovsky,A.G. and Polyakov,K.M. (1990) Trends Biochem. Sci., 15, 158–162.[CrossRef][ISI][Medline]

Shirley,B.A., Stanssens,P., Steyaert,J. and Pace,C.N. (1989) J. Biol. Chem., 264, 11621–11625.[Abstract/Free Full Text]

Steyaert,J. (1997) Eur. J. Biochem., 247, 1–11.[Abstract]

Steyaert,J. and Wyns,L. (1993) J. Mol. Biol., 229, 770–781.[CrossRef][ISI][Medline]

Steyaert,J., Opsomer,C., Wyns,L. and Stanssens, P. (1991) Biochemistry, 30, 494–499.[ISI][Medline]

Strub,K., Moss,J. and Walter,P. (1991) Mol. Cell. Biol., 11, 3949–3959.[ISI][Medline]

Szewczak,A.A., Chan,Y.L., Moore,P.B. and Wool,I.G. (1991) Biochimie, 73, 871–877.[CrossRef][ISI][Medline]

Wang,N., Yamanaka,K. and Inouye,M. (2000) Mol. Microbiol., 38, 526–534.[CrossRef][ISI][Medline]

Wool,I.G., Gluck,A. and Endo,Y. (1992) Trends Biochem. Sci., 17, 266–269.[CrossRef][ISI][Medline]

Yang,X. and Moffat,K. (1996) Structure, 4, 837–852.[ISI][Medline]

Zhong,M. and Kallenbach,N.R. (1994) J. Biomol. Struct. Dyn., 11, 901–911.[ISI][Medline]

Received May 28, 2002; revised September 26, 2002; accepted October 10, 2002.





This Article
Abstract
FREE Full Text (PDF)
Alert me when this article is cited
Alert me if a correction is posted
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Add to My Personal Archive
Download to citation manager
Search for citing articles in:
ISI Web of Science (1)
Request Permissions
Google Scholar
Articles by Chen, D.-T.
Articles by Lin, A.
PubMed
PubMed Citation
Articles by Chen, D.-T.
Articles by Lin, A.