RNase

3.1.3 Regulation of RNase II Expression

RNase II is a monomer of 72 kDa and 644 amino acids (aa), encoded by the rnb gene from E. coli. The rnb gene is transcribed by two promotors, P1 and P2, and terminates in a Rho-independent terminator 10 nts downstream of the rnb stop codon.160 Unless PNPase is absent, RNase II is not an essential enzyme in E. coli,61 but no other RNase can substitute for RNase II to maintain cell survival during nutritional starvation.161 RNase II expression is controlled at the transcriptional and post-transcriptional levels. PNPase cleaves and degrades rnb mRNA, modulating its expression.61 The endonucleases RNase III and RNase E perform an indirect control of RNase II levels. In a mutant strain for RNase III, the pnp mRNA is not properly cleaved, affecting the levels of PNPase and, therefore, the levels of RNase II. In a strain mutant for RNase E, there is an increase in both levels of rnb mRNA and RNase II itself.162 Additionally, it was found that the protein Gmr, which possesses a PAS domain that functions as an environmental sensor, controls the stability of RNase II. It was observed that RNase II is twice more stable in the absence of gmr when compared to the wild-type.163 However, the molecular mechanism underlying this regulation remains mostly elusive.

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Ribonucleases - Part A

Hiroshi Yoshida, in Methods in Enzymology, 2001

Comments

RNase F1 was purified almost quantitatively in four steps, the key step being the affinity chromatography (see Table I). Most impurities flowed through the affinity column, whereas RNase F1 was tightly adsorbed to and eluted specifically from the column by a low concentration of 2′(3′)-GMP. In this particular experiment, the amount of RNase F1 contained in the sample exceeded the capacity of the column and a part of the enzyme appeared in the wash. Even so, however, the unadsorbed RNase F1 was sufficiently retarded to be separated from major impurities. Although most of the brown pigment flowed through the column, a part of it was adsorbed so that the whole column became brown after the chromatography. Therefore, the partially purified preparation from this step contained a small amount of the pigment, which was completely eliminated at the next step, the DEAE-cellulose column chromatography. The affinity column was reusable several times after being washed successively with 5 column volumes each of 0.1 M sodium borate buffer–1 M NaCl (pH 9.0), distilled water, and 0.1 M sodium acetate buffer–1 M NaCl (pH 4.0).

Table I. Purification of RNase F1

ActivityProteinSpecific activityPurificationPurification stepTotal (kU)Yield (%)(A280 U)(kU/A280 U)(–fold)Extraction431010098400.4381Batchwise treatment with DEAE-cellulose473011015203.117.1Affinity chromatography42309812135.079.9DEAE-cellulose column chromatography448010410243.9100

Most enzymes of RNase T1 family are secreted into the fungal culture medium. Only RNase Pol was isolated from a mushroom. Therefore, culture conditions are an important factor for enrichment of an RNase. Although a sufficiently enriched enzyme source of RNase T1 is commercially available (Takadiastase, Sankyo Pharmaceutical Co.), this is an exception. In most cases, purification starts from the culture of a microbe. A search for conditions that favor RNase production may bring a much more enriched source. However, it is not predictable which conditions are suitable for RNase production. For example, Arima et al.23 reported induction of RNase production by Ustilago sphaerogena using RNA. However, addition of RNA was inhibitory for production of RNase F1 by Fusarium moniliforme.24 Instead, yeast extract at a very narrow concentration range (0.1–0.3%) was effective for the production, though it was not clear what component(s) in yeast extract caused the induction. The starting material described above was one order of magnitude richer in RNase F1 than that previously reported.20

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Stability and Stabilization of Biocatalysts

G. Graziano, ... G. Barone, in Progress in Biotechnology, 1998

Ribonuclease P2 from the thermoacidophilic archaebacterium Sulfolobus solfataricus is a small globular protein with a known three-dimensional structure. Inspection of the structure reveals that Phe31 is a member of the aromatic cluster forming the protein hydrophobic core, whereas Trp23 is located on the protein surface and its side-chain is exposed to the solvent. The thermodynamic consequences of the substitution of these two residues in ribonuclease P2 have been investigated by comparing the temperature-induced denaturation of P2 with that of three mutants: F31A-P2, F31Y-P2 and W23A-P2.

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RNAases

E.P. Murchison, in Brenner's Encyclopedia of Genetics (Second Edition), 2013

Enzymology

The majority of RNase enzymes are hydrolytic, catalyzing the hydrolysis of phosphodiester bonds in RNA. The RNase PH family, however, is phosphorolytic, using inorganic phosphate as a reaction substrate for RNA cleavage. RNase enzymes are composed of proteins, some of which function as dimers or as part of oligomeric complexes. RNase P enzymes are ribonucleoproteins whose catalytic activity involves both RNA and protein components. Some RNase enzymes, such as RNase H, use cofactors for catalysis. RNase activity is extremely stable, and can be found in all biological tissues.

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Quantum Leaps in Biochemistry

R.A. Dwek, in Foundations of Modern Biochemistry, 1996

RNase

RNase is an example of a protein which exists in vivo in both nonglycosylated and glycosylated forms, A and B, respectively. RNase B has a single N-linked glycosylation site at Asn-34 and is one of the simplest glycoproteins. Considerable interest has been focused on the differences in the biological functions and properties between RNase A and B (Rudd et al., 1994c). In bovine pancreatic RNase B, biosynthetic processing of the sugar (see below) is halted at the oligomannose stage giving rise to Man5–9GlcNAc2 glycoforms.

Despite the existence of a well-resolved X-ray crystal structure of RNase B, the poor definition of the electron density associated with the oligosaccharide has prohibited any determination of the sugar conformation (Williams et al., 1987). While NMR spectroscopy has been widely applied in the conformational analysis of proteins, including RNase A (Rico et al., 1989, 1991; Robertson et al., 1989: Santoro et al., 1993), and RNase B (Joao et al., 1992), unambiguous conformational determinations of oligosaccharides are less common because of a characteristic paucity of nuclear Overhauser effects (NOEs) between sugar residues. A computer simulation of the dynamic properties of the oligosaccharide offers an alternative approach to the conformational analysis.

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Ribonucleases - Part B

Ti Cai, Mark E. Schmitt, in Methods in Enzymology, 2001

Publisher Summary

Ribonuclease (RNase) MRP is a ribonucleoprotein endoribonuclease that cleaves an RNA sequence in a site-specific manner. The enzyme has two known RNA substrates: one in the mitochondria, where it processes a primer RNA for the initiation of DNA replication, and a second in the nucleus, where it processes the rRNA precursor to help generate the 5.8S rRNA. RNase MRP has been isolated from various organisms including human, mouse, rat, cow, Xenopus, yeast, Arabidopsis, and tobacco. The RNA component of the enzyme was found to be structurally and evolutionally related to RNase P RNA, the ribonucleoprotein endoribonuclease that processes the 5′ end of tRNA, and several protein components are shared between the two enzyme complexes. The data suggest that the enzymes, while ancient, have been highly conserved over time to perform essential cellular RNA-processing functions, both in the nucleus and the mitochondria. This chapter describes methodologies for testing various yeast mutants for endoribonuclease activity both in vivo and in vitro. In addition, a method for testing for assembly of the individual RNase MRP constituents with the complex is described.

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The Role of the 3′ End in mRNA Stability and Decay

Christopher F. Higgins, ... Elisabeth A. Mudd, in Control of Messenger RNA Stability, 1993

B Ribonuclease II

RNase II is similar to PNPase in many respects (Nossal and Singer, 1968; Singer and Tolbert, 1965; Gupta et al., 1977). It too is a processive, 3′–5′ exoribonuclease whose activity is sequence-independent although sensitive to RNA secondary structure. If anything, the degradative activity of RNasell is impeded by RNA secondary structure to an extent greater than that of PNPase (Nossal and Singer, 1968; Gupta et al., 1977; McLaren et al., 1991). The rnb gene, which encodes the 80-kDa RNase II polypeptide, is located at 28 min on the E. coli chromosome (Donovan and Kushner, 1983) and has recently been sequenced (Zilhao et al., 1993). RNaseII differs from PNPase in that it is hydrolytic rather than phosphorolytic and requires potassium ions for full activity.

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Ribonucleases - Part A

Marc Ribó, ... Maria Vilanova, in Methods in Enzymology, 2001

Determination of Steady-State Kinetic Parameters

Ribonuclease cytotoxicity is dependent on its catalytic activity. Thus structure–function studies require appropiate activity measurements. To this end, spectrophotometric assays according to the methodology described by Boix et al.26 can be used to determine the kinetic parameters for the cleavage of poly(C) and the hydrolysis of cytidine 2′,3′-cyclic monophosphate (C > p) by wild-type and mutant HP-RNases. For C > p, the concentration of enzyme is 0.1 μM, the initial concentration of C > p ranges from 0.1 to 5.5 mM, and the activity is measured by recording the increase in absorbance at 296 nm. For assays with poly(C), the concentration of enzyme is 5 nM, the initial concentration of poly(C) ranges from 0.1 to 2.5 mg/ml, and the decrease in absorbance at 294 nm is monitored. Both assays are carried out at 25° in 0.2 M sodium acetate buffer, pH 5.5, using 1 cm path length quartz cells for C > p and 0.2 cm path length quartz cells for poly(C). Steady-state kinetic parameters are obtained by nonlinear regression analysis using the program ENZFITTER.27 The values in Table I are the average of three determinations, with a standard error of less than 10%. Whithin the error of the assay method, no significative differences were found between the HP-RNase variants and the wild-type enzyme (Table I).

Table I. Kinetic and Thermostability Properties of HP-RNase Variants and RNase Aa

C > p substratePoly(C) substrateEnzymeKm (mM)kcat (min−1)Km (mg/ml)(Vmax/[E0])relb (%)T1/2 (°C)PM51.8046.60.44210056.3 ± 1PM71.9355.50.35082.456.1 ± 1PM81.5748.00.399104.455.4 ± 1PM91.0861.60.22564.351.4 ± 1RNase A0.8187.10.43084.758.0 ± 1

aThe rate of cleavage of the high molecular mass substrate poly(C) and the hydrolysis of the low molecular mass substrate C > p were assayed at 25° in 0.2 M sodium acetate, pH 5.5.b[E0]: Total concentration of enzyme in the assay. PM5 is taken as reference.

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Ribonucleases - Part A

L. Aravind, Eugene V. Koonin, in Methods in Enzymology, 2001

α+β Ribonucleases

Ribonuclease PH Fold. The RNase PH superfamily64 defines a compound fold comprised of two distinct substructures.65 The N-terminal subdomain is clearly derived from the ribosomal protein S5 fold that is also seen in other proteins involved in RNA metabolism and translation, such as bacterial RNase P protein component and the elongation factor G domain IV.66,67 In the RNase PH fold, a predicted β-hairpin has been added to the S5 core as an N-terminal extension. This subdomain is packed against another subdomain with a βbα βbα a structure, with the β sheet of this subdomain interacting with the two α helices of the first subdomain. Based on the sequence conservation between the most divergent members of this family, we predict that the active site of these enzymes includes a nearly universal aspartate at the end of the second strand in the second subdomain of the RNase PH domain. A highly conserved arginine located near the N terminus of the first strand in subdomain 1 is in spatial proximity to this aspartate, and is likely to be an important part of the active site. In the same region, also in spatial proximity with the above-mentioned residues, a motif typically containing a RXD signature is conserved in many domains of this superfamily (LA, unpublished observations). These are also likely to be important in stabilizing the active site region. Thus it appears that the RNase PH fold has evolved through elaboration of an S5-like RNA-binding domain into a catalytic domain through extension of a sheet and addition of a subdomain. The members of this superfamily are involved in 3′ processing of RNAs in all the three domains of life and are major components of the exosome in eukaryotes and probably in archaea.36,40,58

Polynucleotide phosphorylases comprise one of the principal families of this superfamily, which is present largely in bacteria, although some eukaryotes, such as Drosophila and Arabidopsis, encode members of this family that probably have been acquired through horizontal gene transfer. This family is characterized by a duplication of the RNase PH domain and fusion to C-terminal KH and S1 RNA-binding domains.64 The duplication of the RNase PH domain results in a quasi-threefold symmetry in these proteins, with only the C-terminal RNase PH domain possessing nuclease activity.65 The typical RNase PH that appears to consist of the catalytic domain alone represents another bacteria-specific family. The RPR45p and Ski6p families are conserved in eukaryotes and archaea and include the nuclease subunits of the exosome.40,58 Eukaryotes additionally have a specific family of exosomal RNase PH: the RPR46 family, that probably evolved through duplication of one of the above archaeoeukaryotic families. There are multiple members of the Ski6p family in C. elegans, one of which contains a catalytic RNase PH domain fused to an uncharacterized domain that is bound to the C terminus of the helicase domain in the RecQ subfamily of helicases.45 The demonstration of the involvement of a RecQ-like helicase in posttranscriptional gene silencing68 suggests that some of these nucleases might contribute to this process in cooperation with RecQ-like helicases.

Metallo-β-Lactamase Fold. Metallo-β-lactamases are a large family of Zndependent hydrolases that contain a characteristic metal binding signature, typically of the form HXHXDH.6 One family from this superfamily includes CPSF-I that is involved in the endonucleolytic cleavage of pre-mRNAs that precedes the addition of the poly(A) tail.69 This family additionally includes the DNA repair enzyme SNM1,70 suggesting that the entire family could be involved in nucleic acid processing functions. The CPSF family consists of four subfamilies. The largest subfamily includes classic CPSF orthologs that are conserved in archaea and eukaryotes and are also seen in certain bacteria. In turn, bacteria have their own, widespread CPSF subfamily that is also represented in several archaea and Arabidopsis. The eukaryotes possess a specific subfamily of inactive CPSF paralogs that are known to form subunits of the CPSF protein complex, probably mediating specific protein–protein interactions.71 The archaea possess a distinct CPSF subfamily in which the metallo-β-lactamase domain is fused to an N-terminal KH domain. In theory, some other, poorly characterized families of the MBL fold also could include nucleases, but there is no direct or circumstantial evidence to predict such activity in any of these proteins.

Barnase Fold. The barnase fold, typified by the barnase superfamily, comprises two known families of secreted RNases from bacteria and fungi, respectively. These proteins have a simple, disulfide bond-stabilized core with four strands with an active site formed by a catalytic triad of E, R, and H residues.72 Their restricted distribution, which is generally characteristic of secreted proteins, suggests that they evolved rather recently as an adaptation for the utilization of environmental polyribonucleotides.

Ribonuclease A Fold. This is another family of secreted nucleases with a very limited distribution; so far, it has been detected only in the vertebrates. The entire fold is stabilized by disulfide bonds, and the active site of these proteins involves a triad of H, K, and H. These proteins appear to be a recent derivation in the vertebrate lineage in relation to pathogen response and environmental polyribonucleotide degradation.73

Helix–Grip Fold. A small number of proteins of the START domain superfamily that possess the Helix–Grip fold74 appear to possess ribonuclease activity.75 These proteins, ribonucleases 1 and 2 from ginseng and their orthologs, belong to the Birch allergen family of START domains that are only known from the plants. These START domains also bind other ligands, such as cytokinins, which is more in line with the ligand-binding activities described for other members of the START superfamily.74 The START domain contains a large ligand-binding channel that could potentially serve as the active site for these proteins. These proteins are known to be induced in response to pathogens in plant tissues and could function as part of a pathogen-specific RNA degradation mechanism.76

DHH Fold. The DHH superfamily of hydrolases is defined by a characteristic DHH signature, which is predicted to contribute to the active site and includes several metal-dependent hydrolases with phosphatase and nuclease activities.5 The domain is predicted to adopt a bipartite α+β fold with the metal-chelating sites located in the N-terminal part. The best-characterized members of this fold are RecJ, a 5′→3′-exonuclease involved in DNA repair in bacteria77 and probably archaea77 and eukaryotic exopolyphosphatase.78 Two families of DHH-fold proteins are predicted to possess RNase activity on the basis of the accessory domains that are fused to their hydrolase domains. The first of these is found only in the archaea and has a unique architecture with the DHH-domain preceded by an N-terminal zinc finger of the DnaJ variety followed by two RNA-binding OB-fold domains, one of the S1 superfamily and one of the N-OB superfamily.79 These proteins could be involved in an archaea-specific 5′→3′ RNA degradation function. Anther DHH family with a potential RNase function is suggested by the fusion of the DHH domain to an N-terminal poly(A) polymerase and CBS domains. These could be bifunctional proteins capable of both degrading bacterial mRNAs and lengthening their poly(A) tails.

Ribonuclease T2 Fold. The RNase T2 superfamily is seen in all crown-group eukaryotes and in some γ-proteobacteria, which suggests a relatively recent horizontal transfer from eukaryotes to bacteria. All these nucleases are secreted and contain a core stabilized by disulfide bonds and an active site with two his-tidines. In most eukaryotes they are likely to be utilized in degradation of extracellular polyribonucleotides, for the acquisition of phosphate or bases. The genes encoding these RNases in plants have undergone multiple duplications and appear to be utilized in other roles such as pathogen response and pollen self-incompatibility.80

KEM1/RAT1 Fold. The KEM1/RAT1 superfamily is defined by a large composite module that is conserved in at least the crown-group eukaryotes. More divergent versions are seen in the earlier-branching eukaryotes, Leishmania and Plasmodium falciparum. The members of this superfamily share a very large conserved region and show a high level of sequence conservation with several invariant polar residues, which hampers prediction of the active site through sequence analysis. Comparisons with the plasmodial form suggest that it might be located in the N-terminal part of the catalytic module. This module probably adopts multiple, distinct substructures of the α+β type. The characterized members of this superfamily function both in the nucleus and in the cytoplasm, in mRNA maturation and turnover, and in 5′-terminal processing of 23S rRNA.81,82 The superfamily consists of two families, the RAT1p family that is represented in all crown-group eukaryotes and the KEM1p family that is restricted to fungi.

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Ribonuclease

Nepomuk Zöllner, Gerd Hobom, in Methods of Enzymatic Analysis, 1965

Sources of Error and Specificity

Apart from ribonuclease, no enzyme is so far known which can catalyse the hydrolysis of cyclic pyrimidine nucleotides. The removal of thermolabile phosphatases, which is necessary when ribonucleic acid is the subtrate for the assay of activity, can be omitted when cyclic pyrimidine nucleotides are used as substrate. Proteins interfere much less in the reaction with cyclic pyrimidine nucleotides than in that with ribonucleic acid. On the other hand the titrimetric method is considerably less sensitive. Ribonuclease can only be determined sufficiently reliably from 2 μg. per reaction mixture, while with the spectrophotometric method 0.25 μg. per reaction mixture can be determined. The precipitation methods permit a reliable measurement of up to 0.003 μg. ribonuclease16), but do not exclude that the action of several enzymes is determined.

*) 1 μequiv. corresponds to 1 ml. 0.001 N NaOH; 1 ml. 0.005 N NaOH corresponds to 5 μequiv.