Sugar non-specific endonucleases

FEMS Microbiology Reviews 25 (2001) 583^613

www.fems-microbiology.org

Sugar non-speci¢c endonucleases E. Srinivasan Rangarajan, Vepatu Shankar * Division of Biochemical Sciences, National Chemical Laboratory, Pune 411 008, India Received 14 May 2001; received in revised form 6 August 2001; accepted 11 August 2001 First published online 26 September 2001

Abstract Sugar non-specific endonucleases are multifunctional enzymes and are widespread in distribution. Apart from nutrition, they have also been implicated in cellular functions like replication, recombination and repair. Their ability to recognize different DNA structures has also been exploited for the determination of nucleic acid structure. Although more than 30 non-specific endonucleases have been isolated to date, very little information is available regarding their structure^function correlations except that of staphylococcal and Serratia nucleases. However, during the past few years, the primary structure, nature of the active site based on sequence homology, and the probable mechanism of action have been postulated for some of the enzymes. This review describes the purification, characteristics, biological role and applications of sugar non-specific endonucleases. ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. Keywords : Endonuclease ; Physicochemical property; Biological role; Application

Contents 1. 2. 3.

4.

5. 6.

7.

8.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Occurrence and localization . . . . . . . . . . . . . . . . . . . Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Viscometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Spectrophotometric methods . . . . . . . . . . . . . . . 3.3. Radioactive measurements . . . . . . . . . . . . . . . . . 3.4. Gel electrophoresis . . . . . . . . . . . . . . . . . . . . . . 3.5. Phosphomonoesterase activity . . . . . . . . . . . . . . Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Agar plate method . . . . . . . . . . . . . . . . . . . . . . 4.2. Zymogram analysis . . . . . . . . . . . . . . . . . . . . . . Puri¢cation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Physical properties . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. Molecular mass and subunit structure . . . . . . . . 6.2. Isoelectric point and glycoprotein nature . . . . . . Catalytic properties . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Optimum pH . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Optimum temperature and temperature stability 7.3. Metal ion requirement . . . . . . . . . . . . . . . . . . . . 7.4. E¡ect of salt concentration . . . . . . . . . . . . . . . . 7.5. E¡ect of denaturants . . . . . . . . . . . . . . . . . . . . . 7.6. Activators and inhibitors . . . . . . . . . . . . . . . . . . Substrate speci¢city . . . . . . . . . . . . . . . . . . . . . . . . .

* Corresponding author. Tel. : +91 (020) 589-3034; Fax: +91 (020) 588-4032.

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . .

584 585 586 586 586 586 587 587 587 587 587 587 588 588 589 590 590 590 591 591 592 592 593

E-mail address: [email protected] (V. Shankar).

0168-6445 / 01 / $20.00 ß 2001 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII: S 0 1 6 8 - 6 4 4 5 ( 0 1 ) 0 0 0 6 9 - 9

FEMSRE 729 5-12-01

584

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

9.

. . . . . . . . . . . . . . . . . .

595 595 595 595 596 596 597 597 598 599 599 599 602 603 604 606 606 607

Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

607

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

607

10. 11. 12. 13. 14. 15. 16.

17. 18. 19.

Mode of action . . . . . . . . . . . . . . . . . . . . . . . 9.1. Endonucleases . . . . . . . . . . . . . . . . . . . . . 9.2. Exonucleases . . . . . . . . . . . . . . . . . . . . . . 9.3. Endo^exonucleases . . . . . . . . . . . . . . . . . Mechanism of action . . . . . . . . . . . . . . . . . . . Action on polynucleotides . . . . . . . . . . . . . . . Action on plasmid DNA . . . . . . . . . . . . . . . . Cleavage preference . . . . . . . . . . . . . . . . . . . . Base speci¢city . . . . . . . . . . . . . . . . . . . . . . . Associated phosphomonoesterase activity . . . . Structure and function . . . . . . . . . . . . . . . . . . 16.1. Active site . . . . . . . . . . . . . . . . . . . . . . . 16.2. Coordination and function of metal ions 16.3. Enzyme^substrate interactions . . . . . . . . 16.4. Water-assisted metal ion catalysis . . . . . Biological role . . . . . . . . . . . . . . . . . . . . . . . . Applications . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Nucleic acids act as carriers of genetic information. For the e¡ective transfer of genetic information from one generation to another, they have to undergo processes such as replication, recombination and repair. All living organisms contain a set of enzymes, namely nucleases, which hydrolyze phosphodiester linkages in nucleic acids. The involvement of enzymes in the break down of nucleic acids was ¢rst observed by Araki [1] and the term `nucleases' was coined by Iwano¡ [2]. Kunitz [3], in 1940, described two classes of nucleases based on their sugar speci¢city. Subsequently, di¡erent schemes of classi¢cation were proposed in an attempt to overcome the shortcomings of the earlier ones [4^6]. However, with the discovery of newer nucleases and multifunctional enzymes, such as micrococcal nuclease [7] and snake venom phosphodiesterase [8], the classi¢cation of Kunitz [3,4] was found to be inadequate. In order to overcome these inadequacies Bernard [9] and Laskowski [5,10] came up with consensus criteria for the classi¢cation of nucleases on the basis of : 1. the nature of substrate hydrolyzed (DNA, RNA); 2. the type of nucleolytic attack (exonuclease or endonuclease) ; 3. the nature of the hydrolytic products formed (i.e. mono- or oligonucleotides terminating in a 3P- or a 5P-phosphate); and 4. the nature of the bond hydrolyzed. Though the above classi¢cation could accommodate enzymes with exceptional properties into a particular group,

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . .

discrepancies still existed as the complex nature of the catalytic activities of di¡erent nucleases was unraveled. Properties related to strand speci¢city, site speci¢city and sequence speci¢city did not ¢nd a place in the above classi¢cation scheme. Additionally, nucleases from Neurospora crassa [11] and BAL 31 nuclease from Alteromonas espejiana [12] exhibited mixed activities, depending on the substrate, making it di¤cult for them to be classi¢ed under a particular scheme. Taking all these aspects into consideration, Linn [13] formulated the most acceptable system of classi¢cation of nucleases (Fig. 1). Nucleases are ubiquitous in distribution and among them sugar non-speci¢c endo- and exonucleases have been implicated in cellular functions like replication, recombination and repair. Their ability to recognize di¡erent DNA structures has made them important analytical tools for the determination of nucleic acid structures [14^ 16], mapping mutations [17] and studying the interactions of DNA with various intercalating agents [18,19]. These enzymes are characterized by their ability to hydrolyze both DNA and RNA. Fraser and Low [20] further subdivided the major sugar non-speci¢c nucleases into three distinct but distantly related groups viz.: 1. secreted fungal single-strand-speci¢c endonucleases ; 2. protease-sensitive multifunctional endo^exonucleases; and 3. mitochondrial nucleases that are closely related to bacterial Serratia marcescens nuclease. Although more than 30 sugar non-speci¢c endonucleases have now been isolated (Table 1), only staphylococcal nuclease [55^57] and S. marcescens nuclease [58]

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

585

Fig. 1. Schematic representation of the classi¢cation of nucleases. Based on the tabulation of nucleases by Linn [13].

have been extensively characterized. However, the past few years have witnessed signi¢cant progress in the molecular enzymology of some of these enzymes. The present review gives a comprehensive account of sugar non-speci¢c endonucleases with respect to their occurrence, puri¢cation, physicochemical properties, biological role and applications. 2. Occurrence and localization It is well known that nucleases play an important role in four Rs, i.e. replication, recombination, restriction and repair. Hence, every living organism must produce at least one type of nuclease. Sugar non-speci¢c endonucleases have been isolated from a wide variety of sources (Table 1). The majority of these enzymes are intracellular but enzymes from Staphylococcus aureus [7], A. espejiana [12], S. marcescens [22], Bacillus subtilis [25], Vibrio sp. [26], Lysobacter enzymogens [27], Saccharomyces cerevisiae [30], Anabaena sp. PCC 7120 [33], Syncephalastrum racemosum [34], Cunninghamella echinulata [38], Streptomyces antibioticus [39] and Rhizopus stolonifer [40] are extracel-

lular in nature. Endonucleases of mitochondrial origin have been reported from N. crassa [11,24], S. cerevisiae [28,29,31] and Aspergillus nidulans [32]. In the cases of N. crassa [24] and A. nidulans [32] their presence has been shown in vacuoles, conidia, mycelia and nuclei. Additionally, in N. crassa, the mitochondrial nuclease has been shown to occur as membrane-bound as well as soluble forms in approximately equal proportions [59]. In plants, sugar non-speci¢c endonucleases have been isolated from tea leaves [42], pollen grains of tobacco [43] and Petunia hybrida [45], wheat chloroplasts and its organelles [46], rye germ ribosomes [47,48] and barley microspores [49]. The presence of these enzymes has also been shown in the lumen of the endoplasmic reticulum, in the Golgi apparatus, in protein bodies and vacuoles of barley aleurone layers [60]. In the kinetoplastid parasite, Leishmania sp., the localization of the endonuclease has been tentatively assigned to nuclei and kinetoplasts [37]. In animals, they have been isolated from various organelles, for example, the hepatopancreas of shrimp [52], rat liver nuclei [50], Bos taurus mitochondria [61] and embryos of Drosophila melanogaster [54].

FEMSRE 729 5-12-01

586

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

decrease indicates an exo mode of action. However, an endo^exonuclease can produce signi¢cant changes in the hyperchromicity as well as a drop in the viscosity.

3. Assay 3.1. Viscometry This method is based on the measurement of decrease in viscosity of the nucleic acid samples following the action of nucleases [62]. 3.2. Spectrophotometric methods The increase in the amount of the acid soluble nucleotides (deoxyribo- or ribonucleotides) produced following the hydrolysis of DNA/RNA is measured at 260 nm [4,63]. A unit of the enzyme is de¢ned on the basis of Wmol of acid soluble nucleotides liberated [64] or Wg of RNA or DNA digested [65]. Viscometric and hyperchromicity measurements can also provide an insight into the mode of action of the enzyme. A sudden drop in the viscosity of the DNA solution without a signi¢cant increase in the hyperchromicity suggests an endonucleolytic cleavage, whereas a gradual

3.3. Radioactive measurements The sensitivity of the assay can be enhanced by the use of radiolabelled substrates, where the increase in the acid soluble radioactivity or decrease in the acid insoluble radioactivity following the action of nuclease is measured. A unit of the enzyme is de¢ned as the amount of enzyme required to render 1 nmol of labelled substrate acid soluble, under the assay conditions [66]. Another rapid and sensitive assay that measures endonucleolytic activity on DNA utilizes the fact that nitrocellulose membrane can retain only large fragments of denatured DNA. In this procedure, following enzyme action, radiolabelled denatured DNA is passed through nitrocellulose ¢lters and the decrease in the retention of radioactivity on the nitrocellulose membrane is measured [67].

Table 1 Sugar non-speci¢c endonucleases Enzyme Microbes Staphylococcal nuclease Azotobacter nuclease Sm nuclease Neurospora nuclease Bacillus nucleases BAL 31 nuclease Marine bacterium nuclease Lysobacter nuclease Yeast nuclease Aspergillus nuclease Anabaena nuclease Sr nuclease Schizosaccharomyces nuclease Streptococcus MF Leishmania nuclease C1 nuclease Streptomyces nucleases Nuclease Rsn Plants Potato nuclease Tea leaves nucleases Pollen nuclease Barley nuclease Pollen nuclease Wheat nuclease Rye germ nuclease-I Rye germ nuclease-II Barley microspores nuclease Animals Rat liver nuclease Endonuclease G Shrimp nuclease Drosophila nuclease Drosophila nuclease

Source

Reference

S. aureus A. agilis S. marcescens N. crassa mitochondria B. subtilis A. espejiana Vibrio sp. L. enzymogens S. cerevisiae mitochondria A. nidulans Anabaena sp. PCC 7120 S. racemosum S. pombe mitochondria S. pyogenes Leishmania sp. C. echinulata var. echinulata S. antibioticus R. stolonifer

[7] [21] [22,23] [11,24] [25] [12] [26] [27] [28^31] [32] [33] [34] [35] [36] [37] [38] [39] [40]

Potato tubers Tea leaves Nicotiana tabacum pollen Barley aleurone layers P. hybrida pollen Wheat chloroplasts Rye germ ribosomes Rye germ ribosomes Hordeum vulgare L. uninucleate microspores

[41] [42] [43] [44] [45] [46] [47] [48] [49]

Rat liver nuclei B. taurus mitochondria Shrimp hepatopancreas D. melanogaster D. melanogaster embryos

[50] [51] [52] [53] [54]

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

587

3.4. Gel electrophoresis

5. Puri¢cation

Agarose gel electrophoresis is a convenient and rapid technique for studying the extent and nature of singleor double-strand breaks, the frequency of damage, and the pattern of distribution of breaks in the substrate [68]. Although developed initially as a qualitative technique, it can be augmented and used as a quantitative method by end-labelling the substrate [69] or by densitometric scanning following electrophoresis in agarose gels [70]. A unit of the enzyme is de¢ned as the amount of enzyme required to produce 1 fmol of nicks in the plasmid DNA under the assay conditions. Separation of the cleavage products on polyacrylamide gel electrophoresis (PAGE) followed by autoradiography can provide information regarding the cleavage site [71].

Since the majority of sugar non-speci¢c endonucleases are intracellular, most of the puri¢cation procedures, irrespective of the source, involve steps like lysis of cells, isolation of organelles (by di¡erential centrifugation), concentration of the crude extract by salt precipitation followed by conventional puri¢cation methods, such as ion-exchange chromatography and gel ¢ltration. During the initial puri¢cation steps, one of the primary aims is to get rid of the colored impurities contributed by pigments of the organelles and this is achieved either by extraction with acetone^water mixture (4:1 v/v) [42] or by a simple wash with ammonium chloride followed by high speed centrifugation [47,48]. In most cases apart from sonication [52], grinding with glass beads or sand [11,21,41,42] has also been used for disrupting the cells. In general, ammonium sulfate precipitation is also used as a preliminary puri¢cation step. In the case of membrane-bound enzymes, solubilization is achieved using non-ionic detergents like Triton X-100 or Nonidet P-40 and/or a high salt wash [31,35,47,77]. Moreover, in certain cases, inhibitors like phenylmethylsulfonyl £uoride (PMSF), pepstatin A and leupeptin are used to overcome proteolytic activity [30,31,77]. Although ion-exchangers like DEAE- and CM-cellulose have been widely used for the puri¢cation of these enzymes, phosphocellulose has also been used in certain cases. For example, potato tuber nuclease, despite its net negative charge at pH 7.5, binds to phosphocellulose due to its a¤nity towards phosphate groups in phosphocellulose [41]. In this manner, this support not only acts as a cation-exchanger but also as an a¤nity matrix. Gel ¢ltration has been used as one of the steps for the puri¢cation of nucleases from Vibrio sp. [26], L. enzymogens [27], S. racemosum [34], S. antibioticus [39], tea leaves [42] and rye germ ribosomes [48]. Hydroxylapatite has been widely used for the puri¢cation of several enzymes including nucleases [78]. This adsorbent has been used successfully for the puri¢cation of endonucleases from S. cerevisiae [28,30], shrimp hepatopancreas [52], S. racemosum [34], Leishmania sp. [37] and B. taurus [77]. Endonucleases from S. racemosum [34], C. echinulata [38], rye germ ribosomes [48] and shrimp hepatopancreas [52] have been puri¢ed on hydrophobic matrices like phenyl- and octyl-Sepharose. A¤nity chromatography has been used extensively for the puri¢cation of sugar non-speci¢c endonucleases. Endonucleases from barley aleurone layers [44], S. cerevisiae mitochondria [31], C. echinulata [38] and R. stolonifer [40] were puri¢ed on heparin-agarose and/or reactive blue 2agarose/Cibacron blue-agarose. Other a¤nity adsorbents include deoxythymidine 3P,5P-diphosphate (pdTp)-aminophenyl-Sepharose [79], NADP-agarose [80] and 5P-AMPagarose [81]. The a¤nity of sugar non-speci¢c endonucleases for single-stranded nucleic acids has been utilized

3.5. Phosphomonoesterase activity Phosphomonoesterase activity, associated with some of the enzymes, is assayed by measuring the inorganic phosphate liberated following the hydrolysis of either 3P-AMP or 5P-AMP. A unit of the enzyme is de¢ned on the basis of Wmol of inorganic phosphate liberated [72]. 4. Detection 4.1. Agar plate method For qualitative detection of nucleases, the nucleic acid (DNA/RNA) is incorporated into the growth medium, in agar plates, along with a dye such as methyl green. The culture to be tested for nuclease production is spotted onto the plate and/or the sample is added in wells on the plates. A clear zone around the growth or the well, after precipitation of the unhydrolyzed methyl green nucleic acid complex with HCl, indicates the presence of nuclease activity [73,74]. 4.2. Zymogram analysis The feasibility of detecting nuclease activity in gels containing nucleic acids was ¢rst demonstrated by Boyd and Mitchell [75]. Following electrophoresis/isoelectric focussing, the gels are incubated in appropriate bu¡ers for enzyme action and then stained with suitable dyes, such as toluidine blue, acridine orange, methyl green or pyronin Y. A clear band against a colored background indicates the presence of nuclease. Alternatively, nucleases that renature after sodium dodecyl sulfate (SDS) treatment can be separated on SDS^PAGE containing the nucleic acid. After electrophoresis, the digested regions in the gel are detected as clear bands against the £uorescent background of ethidium bromide (EtBr) bound to nucleic acids [76].

FEMSRE 729 5-12-01

588

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

for the puri¢cation of endonucleases of S. cerevisiae [30], A. nidulans [32], D. melanogaster [53] and Leishmania sp. [37] on single-stranded DNA (ssDNA) bound to cellulose. In this case, the chromatographic operations are generally carried out under conditions where the enzyme is either not active or shows very little activity. Modern puri¢cation techniques like FPLC have been utilized successfully for the puri¢cation of sugar non-speci¢c endonucleases from S. antibioticus [39], D. melanogaster embryos [54] and B. taurus [77].

nucleases range from 14 to 140 kDa, but the majority of them are between 28 and 44 kDa (Table 2). The enzymes from the marine bacterium Vibrio sp. [26], yeast [29] and A. espejiana F (fast) form [82] are high molecular mass proteins with Mr values of 100, 140 and 109 kDa, respectively. Nucleases from S. aureus [83], yeast mitochondria [28], rye germ ribosomes [47] and S. antibioticus [39] are comparatively low Mr proteins in the range of 14^20 kDa. Most of the non-speci¢c nucleases consist of a single polypeptide chain, but the enzymes from N. crassa mitochondria [24], S. racemosum [34], B. taurus [77] and S. marcescens [84^86] are dimers made up of two identical subunits of 33, 28, 26 and 26 kDa, respectively. Moreover, the dimeric forms of non-speci¢c endonucleases from S. racemosum [34] and S. marcescens [87] are held together purely by non-covalent interactions and not by disul¢de

6. Physical properties 6.1. Molecular mass and subunit structure The molecular masses (Mr ) of sugar non-speci¢c endoTable 2 Properties of sugar non-speci¢c endonucleases Enzyme

Molecular mass (kDa)

Optimum pH

Optimum temperature (³C)

Metal ion requirement

Reference

Azotobacter nuclease Sm nuclease Neurospora nuclease Neurospora nuclease Bacillus nucleases Streptomyces nucleases

^ 52 ^ 66 ^ 18 34 30 28 76 140 14 52 16.8

7.7 8.0^8.5 6.0^7.5 8.0 9.0 8.0^8.5

^ 35^44 37 ^ ^ ^

[21] [23,58] [11] [24] [25] [39]

7.0^8.5 6.8^8.0 7.0^7.5 6.5^7.0 7.0 7.5 9.0^10.0

50^55 ^ 35 ^ 30^40 50 ^

Mg2‡ , Mn2‡ , Mg2‡ , Mn2‡ , Mg2‡ , Mn2‡ , Mg2‡ , Mn2‡ Ca2‡ Mg2‡ Mg2‡ +Ca2‡ Mg2‡ , Mn2‡ Mg2‡ , Mn2‡ , Mg2‡ , Mn2‡ , Mg2‡ , Mn2‡ Mg2‡ , Mn2‡ Mg2‡ , Mn2‡ Ca2‡

109 85 29 72

8.0^8.8 8.0^8.8 7.5 7.5

^ ^ 35 ^

Mg2‡ , Mg2‡ , Mg2‡ , Mg2‡ ,

Ca2‡ Ca2‡ Mn2‡ , Co2‡ Mn2‡

22^28 100 32 67 56 20 57/62 34 36 29 33 60 30

8.0 8.0 7.0^8.0 7.0 7.0^7.2 5.0, 6.5 5.0, 7.8 5.0 6.0 6.8^7.8 6.5^8.0 5.2 ^

^ 30^40 ^ 40^45 ^ ^ ^ ^ 55 50 ^ ^ ^

Mg2‡ , None Mg2‡ , Mg2‡ , Mg2‡ , None None None None None None None Mg2‡

Mn2‡

35 33 32 44 45 50 50

5.5^6.5 5.5^6.5 7.0^8.5 6.5^7.4 7.5 6^7 7.5^8.0

70 60^70 ^ ^ ^ ^ ^

None None Mg2‡ , Mg2‡ , Mg2‡ , Mg2‡ , Mg2‡ ,

C1 nuclease Aspergillus nuclease Yeast nuclease Yeast nuclease Yeast nuclease Leishmania nuclease Staphylococcal nuclease BAL 31 nuclease F (fast) form S (slow) form Anabaena nuclease Yeast nuclease (in£uenced by RAD52) Lysobacter nuclease Vibrio nuclease Schizosaccharomyces nuclease Nuclease Rsn Sr nuclease Rye germ nuclease-I Rye germ nuclease-II Petunia pollen nuclease Barley nuclease Wheat nuclease Potato nuclease Tobacco pollen nuclease Barley microspore nuclease Tea leaves nuclease A nuclease B Drosophila nuclease Drosophila embryo nuclease Shrimp nuclease Rat liver nuclease Endonuclease G

FEMSRE 729 5-12-01

Co2‡ Co2‡ Co2‡

[38] Zn2‡ [32] Co2‡ , Ca2‡ , Zn2‡ [31] [29] [28] [37] [7] [82]

Mn2‡ Mn2‡ , Co2‡ Mn2‡

Mn2‡ Mn2‡ Mn2‡ , Ca2‡ Mn2‡ , Co2‡ Mn2‡

[33] [30] [27] [26] [35] [40] [34] [47] [48] [45] [44] [46] [41] [43] [49] [42]

[53] [54] [52] [50] [77]

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

linkages. The endonuclease from R. stolonifer is a tetramer and each protomer is made up of two non-identical subunits of Mr 21 and 13 kDa. Since the amino acid composition revealed the absence of cysteine [88], it can be assumed that the subunits are held together by non-covalent interactions. A genetically engineered monomeric variant of S. marcescens nuclease, H184R, was shown to exist as a monomer even at high protein concentrations. This monomeric variant exhibited similar secondary structure, stability towards chemical denaturants, and activity to the wildtype enzyme [86]. Although the monomeric variants were functionally independent, in the presence of low enzyme concentration and high molecular mass DNA the native dimeric form of S. marcescens nuclease was relatively more active than the monomeric form or the heterodimer with one inactive subunit [89]. In S. cerevisiae and N. crassa multiple nucleases having similar catalytic properties, but with subtle di¡erences in their physical properties, have been reported. Three endonucleases with Mr of 14, 38 and 57 kDa have been isolated from yeast mitochondria [28,29,31]. Among them, the 38and 57-kDa enzymes exist as dimers under natural conditions. The majority of the nuclease activity in yeast mitochondria, attributed to the 38-kDa enzyme, is located in the mitochondrial inner membrane. Additionally, the presence of multiple forms of the enzyme was correlated to proteolytic cleavage [31]. S. antibioticus produces two extracellular nucleases of Mr 18 and 34 kDa, exhibiting similar properties. The 18-kDa protein was reported to be formed from a 74-kDa precursor, as evidenced by the cross-reactivity against the anti-18-kDa antibodies. It was also shown that the 34-kDa protein is not formed from the 74-kDa protein and the former is not a precursor of the 18-kDa enzyme [39]. In A. nidulans, the 28-kDa endo^exonuclease is formed due to the proteolytic digestion of the 38-kDa polypeptide via the formation of a 33kDa intermediate. Moreover, crude extracts, obtained from cells grown in the presence of PMSF, showed the presence of an immunoreactive 90-kDa polypeptide which is a putative precursor of the 28-kDa endo^exonuclease [32]. In N. crassa 90% of the intracellular neutral ssDNase activity is associated with endo^exonucleases localized in mitochondria, vacuoles and nuclei. In addition, an unstable and trypsin-activatable form of endo^exonuclease found in cytosol, nuclei and mitochondria gave rise to two nucleases similar to endo^exonucleases reported by Linn and Lehman [90,91] and Fraser et al. [92]. From mitochondria, a strand non-speci¢c endonuclease [11] and a ssDNA binding endo^exonuclease not inhibited by ATP [24] have also been isolated. Fraser et al. [93] demonstrated by immunochemical techniques that all the above enzymes from N. crassa are derived from one or more related large inactive (precursor) polypeptides which undergo proteolysis to produce various active forms of nucleases. This inactive precursor was shown to have a Mr of 93^95 kDa [94]. Additionally, Fraser et al. [93]

589

showed that endo^exonucleases from A. nidulans and S. cerevisiae are immunologically related to N. crassa endo^ exonuclease but not to single-strand-speci¢c nucleases like S1, P1 and mung bean nuclease. Similarly, isoforms of the nucleases were observed in S. aureus [95], B. subtilis [25], A. espejiana [12], S. marcescens kums 3958 [23], tobacco pollen [43], P. hybrida pollen [45] and S. marcescens [96^ 98]. In case of staphylococcal nuclease, Taniuchi and An¢nsen [99] showed that limited proteolysis of the enzyme, in the presence of Ca2‡ and pdTp, produces an active noncovalently bonded derivative, termed nuclease-T, which shows a similar level of activity against DNA and RNA as that of the native enzyme. In contrast to protease-mediated formation of multiple forms, shrimp nuclease exhibited multiple forms in the presence of SDS and 2-mercaptoethanol. The enzyme is a monomer of Mr 45 kDa cross-linked with a large number of disul¢de bridges. It exhibits four di¡erent forms on SDS^PAGE, depending on the treatment of the sample (Fig. 2). The native Form I of the protein (which is acidic in nature) is represented by a structure in which the acidic groups (carboxylate residues) are exposed and the disul¢des are buried. When the protein was subjected to heat treatment with 2-mercaptoethanol, in the presence or absence of SDS, it changed into an inactive Form III (45 kDa, reaction a). Form III could not refold and re-oxidize to form active Form I even after the removal of SDS and 2-mercaptoethanol (reaction aP). However, when Form I was subjected to heat treatment with SDS in the absence of 2-mercaptoethanol, the resulting Form II (39 kDa, reaction b) became catalytically active only after the removal of SDS (reaction bP). The native protein on heat treatment in the absence of SDS became irreversibly denatured (Form IIP, reaction c) which could not refold back to the active Form I (reaction cP). However, the Form IIP could not be converted to Form II, even after treatment with SDS (reaction d). Based on these observations the authors opined that the irreversible denaturation on heating in the absence of SDS is due to the formation of the incorrect inside-out structure, with an exposed hydrophobic domain which has a tendency to aggregate and cannot revert to a catalytically active form [100]. 6.2. Isoelectric point and glycoprotein nature The isoelectric points of the majority of sugar non-speci¢c endonucleases have not been reported, but in the case of some of the well characterized enzymes they are in the range 4.0^9.6. Nucleases from shrimp hepatopancreas [52], A. espejiana [82], R. stolonifer [40], rye germ ribosomes [47], D. melanogaster embryos [54] and S. racemosum [34] are acidic proteins with pI values of 4.06, 4.2, 4.2, 4.8, 4.9 and 5.0, respectively. S. marcescens nuclease and its isoforms are neutral proteins with pI values of 6.8, 7.3, 7.4 and 7.5 [23,97]. However, the endonuclease SM1 from S. marcescens kums 3958 [23] and S. aureus nuclease [101]

FEMSRE 729 5-12-01

590

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

Fig. 2. A scheme for folding and unfolding pathways of shrimp nuclease in the presence and absence of SDS and 2-mercaptoethanol (reprinted from Lin et al. [100], Copyright 1994, with permission from Elsevier Science).

are basic proteins with pI values of 8.1 and 9.6, respectively. Nucleases from barley aleurone layers [102], S. racemosum [52] and rye germ ribosomes [47] are glycoproteins. Rye germ ribosomal nuclease I contains 28% carbohydrate consisting of fucose, mannose and glucosamine [47]. 7. Catalytic properties 7.1. Optimum pH In general, the pH optima of sugar non-speci¢c endonucleases are in the range 5.0^10.0 (Table 2) and most of the enzymes show the same optimum pH for the hydrolysis of both polymeric and monomeric substrates. However, nucleases from A. espejiana [81], N. crassa (mitochondria) [11] and tea leaves [42] show di¡erent pH optima for the hydrolysis of ssDNA (8.8, 6.5^7.5 and 6.0) and doublestranded DNA (dsDNA) (8.0, 5.5^6.5 and 5.5, respectively). On the other hand, 3P-nucleotidase-nuclease from potato tubers showed di¡erent pH optima for the nucleotidase (pH 8.0) and nuclease (pH 6.5^7.5) activities [41]. Similarly, nuclease I from rye germ ribosomes exhibited pH optima of 6.0 and 6.5 for the nuclease and 3P-nucleotidase activities, respectively. Nucleases from wheat chloroplasts [46] and rye germ ribosomes [48] showed optimum

pH values of 7.8 and 5.0 for the hydrolysis of denatured DNA and 6.8 and 7.8 for the hydrolysis of RNA, respectively. However, rye germ ribosomal nuclease II degraded poly(I).poly(C) optimally at pH 8.5 [48]. Interestingly, nucleases (A and B) from tea leaves exhibited di¡erent pH optima for the hydrolysis of di¡erent 3P-mononucleotides (i.e. pH 6.0 for 3P-GMP and 3P-CMP and pH 6.5 for 3PAMP and 3P-UMP), a property associated with the 3Pnucleotidase activity of single-strand-speci¢c nucleases like Le1 and Le3 from Lentinus edodes [103,104] and P1 from Penicillium citrinum [64]. In certain cases, the optimum pH of sugar non-speci¢c endonucleases is in£uenced by the type of metal ions involved. For example, S. racemosum nuclease exhibited optimum pH values of 7.0 and 7.2 in the presence of 5 mM Mg2‡ or Mn2‡ , respectively [34]. However, the DNase activity of staphylococcal nuclease showed an optimum pH of 10.0 or 9.5 in the presence of 1 or 10 mM Ca2‡ , respectively, but under similar conditions the RNase activity was optimal at pH 9.5 or 9.0 [105]. 7.2. Optimum temperature and temperature stability The temperature optima of the majority of sugar nonspeci¢c endonucleases have not been reported but in the cases of some of the well characterized enzymes, they are in the range of 30^70³C (Table 2). Most of them exhibit the same optimum temperature for all the substrates.

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

However, C. echinulata nuclease showed an optimum temperature of 50³C and 55³C for DNA hydrolysis in the presence of Mg2‡ and Mn2‡ , respectively [38]. Nuclease Rsn from R. stolonifer showed an optimum temperature of 40³C for ssDNA hydrolysis in the presence of Mg2‡ , Mn2‡ and Co2‡ and for dsDNA hydrolysis in the presence of Mg2‡ , but it showed a higher optimum temperature (45³C) for dsDNA hydrolysis in the presence of Mn2‡ and Co2‡ [40]. On the contrary, the RNase activity of nuclease Rsn exhibited an optimum temperature of 35³C [106]. The majority of sugar non-speci¢c endonucleases are thermolabile enzymes, but staphylococcal nuclease showed high stability and retained a signi¢cant amount of its activity in the presence of 1 N HCl for 6 h or at 100³C for 20 min. Similarly, tea leaf nucleases were stable at 60³C for 15 min [42], whereas nucleases SM1 and SM2 from S. marcescens kums 3958 retained their activity for 24 h at 25³C [23]. Interestingly, a mitogenic factor of Streptococcus pyogenes exhibiting nuclease activity was resistant to inactivation at 100³C for 10 min and showed a biphasic temperature stability. The enzyme was stable up to 50³C (for 10 min) but lost more than 90% of its activity at 60³C. However, it regained approximately 50% of its original activity at 80³C [36]. 7.3. Metal ion requirement Sugar non-speci¢c endonucleases, with the exception of nucleases from Vibrio sp. [26], potato tubers [41], tea leaves [42], barley aleurone layers [44], P. hybrida [45], wheat chloroplasts [46] and rye germ ribosomes [47,48], are metal requiring enzymes. Although wheat chloroplast nuclease [46] did not require metal ions for its activity, the ssDNase activity showed slight stimulation (20%) in the presence of Mg2‡ . Similarly, the activity of endonuclease from Vibrio sp. was stimulated by Mg2‡ and Ca2‡ [26]. In contrast, nuclease I of rye germ ribosomes [47], pollen nucleases of tobacco [43] and P. hybrida [45] were stimulated by Zn2‡ , while nuclease II from rye germ ribosomes was stimulated by Mn2‡ [48]. Extracellular nucleases (BsIA, Bs-IB and Bs-II) from B. subtilis required Ca2‡ for the hydrolysis of native DNA. However, low rates of hydrolysis of denatured DNA and rRNA were observed in the absence of added metal ion [25]. The action of N. crassa nuclease on dsDNA is dependent on Mg2‡ concentration but its activity on ssDNA is independent of Mg2‡ concentration, though it is stimulated to some extent [107]. Addition of 10 mM Mg2‡ , Ca2‡ or Fe2‡ resulted in a 2.5-fold stimulation of the ssDNase activity of N. crassa enzyme, but it also brought about approximately 40% inhibition of the RNase activity. The selective inhibition of the RNase activity in the presence of metal ions was attributed to the induction of secondary structures in RNA by these metal ions. On the contrary, nuclease Rsn from R. stolonifer required Mg2‡ , Mn2‡ and Co2‡ for its DNase and RNase

591

activities. Although Mg2‡ and Mn2‡ did not a¡ect the ssDNase to dsDNase ratio, the enzyme showed a higher preference for ssDNA in the presence of Co2‡ . Moreover, there was no synergism in the presence of either of the metal ions but a higher ssDNase:dsDNase ratio was observed when Co2‡ was used in combination with Mg2‡ or Mn2‡ [40]. Many of the metal requiring endonucleases show synergism when metal ions are used in combination. For example, nucleases from S. aureus [105], A. espejiana [12], A. nidulans [32], shrimp hepatopancreas [52], mitogenic factor of S. pyogenes [36] and S. antibioticus (18-kDa nuclease) [39] exhibited maximum activity in the presence of Ca2‡ and Mg2‡ . 7.4. E¡ect of salt concentration It has been reported that salt concentration in the reaction mixture can in£uence the activity of sugar non-speci¢c endonucleases. For example, the activity of BAL 31 nuclease is maximum in the range of 0^2 M NaCl and the enzyme shows only 40% of its ssDNase activity in the presence of 4.4 M NaCl [12]. In the case of endonucleases from N. crassa [107] and yeast mitochondria [30], 200 mM NaCl completely inhibited the dsDNase activity but it had only a marginal e¡ect on the ssDNase activity. Similarly, D. melanogaster endonuclease showed 50% inhibition of its dsDNase activity in the presence of 30 mM NaCl but it required 100 mM NaCl to bring about the same level of inhibition of the ssDNase activity [53]. The inhibition of dsDNase activity, in the presence of high salt concentrations, was attributed to the stabilization of the AT-rich regions in dsDNA [108,109]. The ssDNase and RNase activities of yeast mitochondrial nuclease are maximal between 1 and 300 mM KCl while KCl ( s 150 mM) inhibited the dsDNase activity. However, NaCl up to 100 mM did not have any e¡ect on the enzyme activities [31]. Similarly, staphylococcal nuclease [55] and endonuclease G from B. taurus [51] were inhibited at NaCl concentrations above 100 mM. On the contrary, Leishmania endonuclease showed optimal activity in the presence of 100^150 mM NaCl/KCl but subsequent increase in the salt concentration progressively inhibited the enzyme activity [37]. Nucleases from S. antibioticus [39] and yeast mitochondria [28], however, showed maximum activity in the presence of 20^30 mM NaCl and 50 mM KCl, respectively. The dsDNase activity of tobacco pollen nuclease was inhibited in the presence of 200 mM NaCl [43] while Serratia nuclease retained 25% of its activity in the presence of 1 M NaCl [110]. For the mitogenic factor of S. pyogenes concentrations of NaCl/KCl greater than 60 mM brought about 90% inhibition of the nuclease activity [36]. Inhibition in the presence of high concentrations of NaCl/KCl was also observed with endonucleases from rat liver nuclei [50], Anabaena sp. [33], Schizosaccharomyces pombe [35] and R. stolonifer [40,106].

FEMSRE 729 5-12-01

592

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

7.5. E¡ect of denaturants In the presence of 7 M urea, S. marcescens endonuclease showed approximately 2-, 1.7- and 1.5-fold increases in its dsDNase, ssDNase and RNase activities, respectively [111]. However, Azotobacter agilis endonuclease showed a 5^10-fold stimulation of its activity towards poly(A) in the presence of 2 M urea [21]. Gray et al. [12] showed that the S (slow) form of BAL 31 nuclease is active in the presence of 5% (w/v) SDS and can be incubated with the detergent without loss of activity if Ca2‡ and Mg2‡ are present at concentrations of 12.5 mM before the addition of the detergent. Puri¢ed S form BAL 31 nuclease retained 60% of its optimal activity in the presence of 4 M urea. In the case of endonuclease from barley aleurone layers, complete inactivation of the DNase and RNase activities was observed in the presence of 1% (w/v) SDS [102]. Similarly, the DNase activity of nuclease Rsn showed high sensitivity towards denaturants and lost more than 80% of its activity in the presence of low concentrations of SDS (0.02% w/v), guanidine hydrochloride (0.2 M) or urea (3 M) [40]. 7.6. Activators and inhibitors Polyamines such as spermine and spermidine, which bind to double-stranded nucleic acids, also inhibit the ssDNase activity of nucleases. Spermine stimulated the exonuclease activity of BAL 31 nuclease but the cleavage speci¢city was considerably reduced in its presence [112]. The DNase and RNase activities of yeast mitochondrial endonuclease [31] and the RNase activity of rye germ ribosomal nuclease I [47] were stimulated in the presence of 2 and 2.5 mM spermidine, respectively. Moreover, a low concentration (0.1 mM) of polyamines such as putrescine and spermidine inhibited the RNase activity of rye germ ribosomal nuclease I [47]. On the contrary, the DNase and RNase activities of S. aureus nuclease were stimulated by low amounts of putrescine, spermine and spermidine, whereas at high concentrations they inhibited both the activities [113]. However, endonuclease from barley aleurone layers was not a¡ected by putrescine, spermidine and spermine at 1 mM [102]. The ability of polyamines to either activate or inhibit the nuclease activity in a concentration-dependent manner was correlated to the regulation of nucleic acid levels within the cells by controlling the nuclease activity [114]. Low concentrations of EtBr (1^10 Wg ml31 ) brought about marginal stimulation of the dsDNase activity of yeast mitochondrial endonuclease but it had no e¡ect on the RNase activity [31]. Although 10 WM EtBr did not a¡ect the ssDNase activity of yeast mitochondrial endonuclease, slight inhibition (30%) of the dsDNase activity was observed with 2 WM EtBr [28]. Endonucleases from D. melanogaster [53] and S. antibioticus [39] showed optimal activity in the presence of either dithiothreitol (DTT) or 2-mercaptoethanol. Al-

though N. crassa nuclease [11] and rye germ ribosomal nuclease I [47] did not show an obligate requirement for thiol reagents for their activity, considerable stimulation of the activity was observed in the presence of DTT and 2mercaptoethanol. In contrast, DTT and 2-mercaptoethanol brought about signi¢cant inhibition of both the DNase and RNase activities of wheat chloroplast nuclease [46]. Ten mM DTT inhibited barley aleurone layer nuclease whereas 2-mercaptoethanol, at the same concentration, had no e¡ect on the enzyme activity [102]. However, sulfhydryl reagents had no e¡ect on endonucleases from S. marcescens kums 3958 [23], yeast mitochondria [31] and shrimp hepatopancreas [52]. Interestingly, 2-mercaptoethanol had no e¡ect on the activity of S. marcescens endonuclease [110], though disul¢de bonds are essential for the enzyme activity [87]. Subsequently, Filiminova et al. [111,115] demonstrated that substantial inactivation of the enzyme activity occurred when a high concentration of 2-mercaptoethanol (640 mM) was used in the presence of 2 M urea. The inability of 2-mercaptoethanol to inactivate the native enzyme was correlated to the masking of the sulfhydryl groups in the native conformation. The majority of the sugar non-speci¢c endonucleases are metal requiring enzymes and hence they are strongly inhibited by metal chelators like EDTA and EGTA. In most cases, the inactivation can be readily reversed by the addition of divalent cations. The ssDNase activity of wheat chloroplast nuclease was strongly inhibited by EDTA but it had no signi¢cant e¡ect on its RNase activity [46]. Endonucleases from rat liver nuclei [50] and S. marcescens kums 3958 [23] were inhibited by pyrophosphate, whereas the enzyme from tobacco pollen was inhibited by inorganic phosphate [43]. While inorganic phosphate signi¢cantly inhibited the DNase and RNase activities of B. subtilis nuclease [25], it only inhibited the RNase activity of rye germ ribosomal nuclease I [47]. On the other hand, inorganic phosphate had no e¡ect on staphylococcal nuclease [105]. However, inorganic phosphate and pyrophosphate inhibited both the DNase and RNase activities of nuclease Rsn [40,106]. Divalent cations like Mn2‡ , Co2‡ and Zn2‡ inhibited nucleases from potato tubers [41] and B. subtilis [25] while wheat chloroplast nuclease was inhibited by Cu2‡ , Co2‡ and Zn2‡ [46]. Nucleases from S. marcescens kums 3958 [23], shrimp hepatopancreas [52] and D. melanogaster [53] were inhibited by Ca2‡ . Though Zn2‡ preferentially inhibited the exonuclease activity on dsDNA of N. crassa mitochondrial nuclease [107] and the RNase activity of rye germ ribosomal nuclease I [47], it inhibited both the DNase and RNase activities of S. aureus [105] and S. antibioticus [39] nucleases. Additionally, Hg2‡ and Cd2‡ inhibited the DNase and RNase activities of staphylococcal nuclease, whereas Ba2‡ and Sr2‡ inhibited only its RNase activity [105]. The inhibition of the RNase activity, in the presence of Mg2‡ and Ca2‡ , was also observed with rye germ ribosomal nuclease I [47]. While Mn2‡ inhibited

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

the endonuclease activity of barley microspores nuclease [49], tea leaf nucleases (A and B) were inhibited by Mg2‡ , Mn2‡ , Co2‡ , Cu2‡ and Fe3‡ [42]. Interestingly, in case of tea leaf nucleases, Zn2‡ did not inhibit the dsDNase activity but showed considerable inhibition of the ssDNase, RNase and 3P-nucleotidase activities. However, Mg2‡ selectively inhibited the DNase and RNase activities with no e¡ect on the 3P-nucleotidase activity [43]. In case of nuclease Rsn, Zn2‡ , Cu2‡ and Hg2‡ inhibited the DNase activity [40] but Cu2‡ and Hg2‡ had no e¡ect on the RNase activity [106]. Nucleotides and their analogues are potent inhibitors of sugar non-speci¢c endonucleases. For example, staphylococcal nuclease was inhibited by pdTp, 5P-dAMP and 5PdTMP. While 5P-dCMP and 5P-dGMP inhibited the RNase activity, the DNase activity was not a¡ected. Moreover, among ribonucleotides, only 5P-AMP could strongly inhibit both activities of the enzyme [105]. Thymidine 3P,5P-diphosphate and ATP also inhibited nucleases SM1 and SM2 from S. marcescens kums 3958, whereas adenosine had no e¡ect [23]. In contrast, ATP did not have any e¡ect on the activity of yeast mitochondrial endonucleases [30,31] and endonuclease G from B. taurus [51]. In the case of potato tuber nuclease, the 3P-nucleotidase activity was inhibited competitively by 5P-mononucleotides, RNA and poly(A) [116]. S. antibioticus nucleases were inhibited by aurin tricarboxylic acid [39] while heparin, Cibacron blue and aurin tricarboxylic acid inhibited yeast mitochondrial endonuclease [31]. Rat liver endonuclease showed inhibition in the presence of polydextran sulfate [50]. The endo^exonuclease from N. crassa was inhibited by a heat-stable, trypsin-sensitive, cytosolic 24-kDa polypeptide. The protein inhibited the ssDNase activity non-competitively but the dsDNase activity was inhibited competitively. In addition, the inhibitor blocked the formation of site-speci¢c double-strand breaks and nicking of linearized pBR322 DNA. It also inhibited the RNase activity of N. crassa nuclease as well as the immunochemically related nuclease from A. nidulans [117]. Moreover, the antibodies raised against N. crassa endo^exonuclease inhibited all the activities of yeast mitochondrial nucleases [30,31]. The endonuclease NucA from Anabaena sp. was inhibited by its polypeptide inhibitor NuiA, while the related Serratia nuclease was not inhibited with a 10-fold excess of the inhibitor [118]. Cleavage of the monomeric substrate deoxythymidine 3P,5P-bis-(p-nitrophenyl phosphate) by NucA, however, was not inhibited by NuiA suggesting that small molecules gain access to the active site of NucA in the enzyme^inhibitor complex under conditions where cleavage of DNA is completely inhibited [119]. Similarly, the extracellular nuclease (Nuc) from S. marcescens was inhibited by the signal peptide obtained from the N-terminal portion of Nuc [120]. Unlike single-strand-speci¢c nucleases, namely S1 nuclease from Aspergillus oryzae [121], nuclease L from Ustilago

593

maydis [122] and nuclease Bh1 from Basidiobolus haptosporus [123], auto-retardation due to end-product inhibition has not been reported in sugar non-speci¢c endonucleases. On the contrary, staphylococcal nuclease showed auto-acceleration during the hydrolysis of denatured DNA, native DNA and RNA. The phenomenon, best observed with polynucleotides intermediate in size between nucleic acids and short oligonucleotides, was attributed to the presence of substantial `breathing spaces' in the intermediate size substrates [124]. 8. Substrate speci¢city Sugar non-speci¢c nucleases are multifunctional enzymes and exhibit both endo- and exonuclease activities (Table 3). Moreover, endonucleases from potato tubers [41], tea leaves [42], barley aleurone layers [44] and rye germ ribosomes [47] exhibit 3P-phosphomonoesterase activity. Sugar non-speci¢c endonucleases act on DNA, RNA and 3P-mononucleotides but the rate of hydrolysis of these substrates varies depending on the source of the enzyme. Thus, potato tuber nuclease showed higher activity on 3PAMP and RNA [41], whereas rye germ ribosomal nuclease I preferred DNA to RNA and 3P-mononucleotides [47]. Nucleases from tea leaves [42] and barley aleurone layers [44] showed preference for RNA in comparison to DNA and 3P-mononucleotides. The substrate speci¢city of potato tuber nuclease falls in the order 3P-AMP = RNA s ssDNA s dsDNA [41] while that of rye germ ribosomal nuclease I is ssDNA s dsDNA s RNA s 3P-AMP [47]. Tea leaf nucleases [42] and barley aleurone nuclease [102] showed substrate preferences in the order of RNA s ssDNA = dsDNA s 3P-AMP. The endonucleases from S. marcescens [110,126], Anabaena sp. [33], N. crassa [11] and S. racemosum [34] hydrolyzed ssDNA, dsDNA and RNA at a similar rate while staphylococcal nuclease [105] and nuclease Rsn from R. stolonifer [40] hydrolyzed both DNA and RNA with a slight preference for ssDNA. Similar preferences for ssDNA were also shown by endonuclease M (Endo M) from Leishmania sp. [37] and yeast mitochondrial endonuclease [31]. In addition, Endo M degraded ssRNA rapidly but a RNA :DNA hybrid was resistant to cleavage. With increasing concentrations of Endo M, the unlabelled single-stranded overhang of DNA from the RNA :DNA hybrid was cleaved to give the perfect duplex RNA:DNA hybrid. However, in the presence of a 10-fold excess enzyme, the resulting RNA :DNA hybrid was also cleaved [37]. Similarly, endonuclease G from B. taurus showed RNase H activity in addition to DNase and RNase activities [61]. Interestingly, nucleases from L. enzymogens [27], shrimp hepatopancreas [52], S. racemosum [34] and S. antibioticus [39] showed preference for dsDNA. Although 18and 34-kDa nucleases from S. antibioticus were reported

FEMSRE 729 5-12-01

594

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

Table 3 Substrate speci¢city and mode of action of sugar non-speci¢c endonucleases Enzyme

Substrate

ssDNase : dsDNase ratio

Products

Mode of action

Reference

Azotobacter nuclease Sm nuclease Neurospora nuclease Neurospora nuclease Bacillus nucleases Streptomyces nucleases 18 kDa 34 kDa C1 nuclease Aspergillus nuclease Yeast nuclease Yeast nuclease Yeast nuclease Staphylococcal nuclease BAL 31 nuclease F (fast) form S (slow) form Leishmania nuclease Anabaena nuclease Yeast nuclease (in£uenced by RAD52) Lysobacter nuclease Vibrio nuclease Schizosaccharomyces nuclease Nuclease Rsn Rye germ nuclease-I

DNA, DNA, DNA, DNA, DNA,

RNA RNA RNA RNA RNA

ss v ds ss = ds ss v ds ss v ds 1.3

5P-oligonucleotides 5P-mono-, di-, tri- and tetranucleotides 5P-mono-, di-, tri- and tetranucleotides 5P-di-, tri-, tetra- and pentanucleotides 3P-mononucleotides

endo endo endo endo^exo endo^exo

[125] [23,126] [11] [24] [25] [39]

DNA, DNA, DNA, DNA, DNA, DNA, DNA, DNA,

RNA RNA RNA RNA RNA RNA RNA RNA

ss s ds ss 6 ds ss = ds 5 2 ss v ds ss v ds 2

5P-mono- and dinucleotides 5P-mono- and dinucleotides 5P-oligonucleotides 5P-di-, tri-, tetranucleotides 5P-products 5P-products 5P-mono-, di- and trinucleotides 3P-mononucleotides

endo/exo endo/exo endo endo endo^exo endo endo endo

DNA, DNA, DNA, DNA, DNA,

RNA RNA RNA RNA RNA

3 33 2 ss = ds 8

5P-mononucleotides 5P-mononucleotides 5P-mononucleotides and oligonucleotides ^ 5P-mononucleotides and oligonucleotides

endo^exo endo^exo endo endo endo

[37] [33] [30]

DNA, RNA DNA, RNA DNA, RNA DNA, RNA DNA, RNA, 3Pmonoribonucleotides DNA, RNA DNA, RNA

ss v ds ss v ds ss v ds 2 1.14

endo endo endo endo endo

[27] [26] [35] [40] [47]

endo endo

[45] [46]

endo endo

[48] [102] [116]

Petunia pollen nuclease Wheat nuclease Rye germ nuclease-II Barley nuclease Potato nuclease Tea leaves nuclease A nuclease B Tobacco pollen nuclease Barley microspore nuclease Drosophila nuclease Drosophila embryo nuclease Shrimp nuclease Sr nuclease Rat liver nuclease Endonuclease G

DNA, RNA DNA, RNA, 3Pmonoribonucleotides DNA, RNA, 3Pmonoribonucleotides

1.14 ss v ds

5P-products 5P-products 5P-products 5P-di-, tri- and tetranucleotides 5P-oligoribonucleotides, 3Poligodeoxyribonucleotides 5P-oligonucleotides 3P-oligoribonucleotides, 5Poligodeoxyribonucleotides 5P-oligonucleotides 5P-oligonucleotides

3

5P-mononucleotides

endo

DNA, RNA, 3Pmonoribonucleotides DNA, RNA, 3Pmonoribonucleotides DNA, RNA DNA, RNA DNA, RNA DNA, RNA DNA, RNA DNA, RNA DNA, RNA DNA, RNA (DNA:RNA)

1

5P-mononucleotides and oligonucleotides

endo

1

5P-mononucleotides and oligonucleotides

endo

3 ss v ds ss = ds ds s ss ds s ss ds s ss 7 ds v ss

5P-mono- and oligonucleotides 5P-products 5P-mono- and oligonucleotides 5P-products ^ ^ s 5P-tetranucleotides 5P-products

endo^exo endo endo^exo endo endo endo endo endo

4 15

[38] [32] [31] [29] [28] [7] [82]

[42]

to show preference for dsDNA [39], subsequently it was shown that 18-kDa nuclease prefers ssDNA (J. Sanchez, unpublished results). Synthetic substrates like deoxythymidine 3P,5P-bis-(p-nitrophenyl phosphate) have been employed for the elucidation of the kinetics and mechanisms of a number of phosphodiesterases [127,128]. Exonucleases, e.g. snake venom phosphodiesterase [129] and spleen phosphodiesterase [130], and endonucleases, e.g. pancreatic DNase [131], could cleave the synthetic substrate. Cuatrecasas et al.

[43] [47] [53] [54] [52] [34] [50] [77]

[132] studied the hydrolysis of various p-nitrophenyl ester derivatives of deoxythymidine 5P-phosphate by staphylococcal nuclease and demonstrated that p-aminophenyl, pnitrophenyl or methyl derivatives are hydrolyzed rapidly. Based on these experiments it was proposed that staphylococcal nuclease requires R-pdT-RP as the basic structural unit to be recognized as a substrate and the RP group at C-3P signi¢cantly in£uences the binding of the substrate or the inhibitor. Additionally, in the absence of R (i.e. free 5P-phosphate), the activity is completely inhibited as evi-

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

595

ing endonucleases, and nucleases from S. marcescens, N. crassa and S. aureus belong to this group. However, except for S. marcescens, N. crassa and staphylococcal nucleases, all the other enzymes mentioned above act mainly on DNA (especially dsDNA) and hence they will not be elaborated in this compilation. 9.2. Exonucleases These enzymes generally require a free terminus for their action and are incapable of hydrolyzing covalently closed circular substrates. However, an exonuclease like snake venom phosphodiesterase is capable of nicking supercoiled DNA [135]. The products of hydrolysis are predominantly mononucleotides and the mode of attack is processive. Spleen phosphodiesterase and A. sydowii nuclease belong to this category of enzymes. Though these enzymes are sugar non-speci¢c, they are strict exonucleases and hence will not be discussed in detail. Fig. 3. Cleavage speci¢cities of di¡erent nucleases (reproduced with permission from Friedho¡ et al. [133]).

denced by strong inhibition with pdTp [105]. Friedho¡ et al. [133] demonstrated that Serratia nuclease hydrolyzes the arti¢cial minimal substrate, deoxythymidine 3P,5P-bis(p-nitrophenyl phosphate), at a lower rate than DNA and RNA. The cleavage sites of staphylococcal nuclease, Serratia nuclease and pancreatic DNase I are shown in Fig. 3. Interestingly, with Serratia nuclease, the presence of phosphate 3P to the bond to be cleaved is essential for hydrolysis [133,134]. Similar observations were made in the case of Anabaena nuclease [119]. 9. Mode of action Although sugar non-speci¢c endonucleases recognize and hydrolyze a wide spectrum of substrates, they primarily cleave the internucleotide phosphodiester linkage. Based on the requirement for a free terminus, these enzymes can be classi¢ed as endonucleases, exonucleases or endo^exonucleases. 9.1. Endonucleases This class of enzymes cleave the internal phosphodiester bonds of nucleic acids with or without free termini. Their ability to convert covalently closed circular DNA to a linear form, either directly or through a nicked circular DNA intermediate, is still considered as one of the criteria for inclusion of these enzymes in this group. They show a distributive mode of action and the products of hydrolysis are mononucleotides and/or oligonucleotides. Restriction endonucleases, structure-speci¢c enzymes like £ap endonuclease, Holliday junction resolvase, sequence-speci¢c hom-

9.3. Endo^exonucleases The enzymes belonging to this category exhibit both exo and endo modes of action. Sugar non-speci¢c nucleases hydrolyze both DNA and RNA, either endonucleolytically or exonucleolytically but some enzymes exhibit di¡erent modes of action on these substrates (Table 3). The majority of these enzymes degrade nucleic acids to oligonucleotides and some mononucleotides predominantly by their endonucleolytic action. Nucleases from N. crassa [24], tobacco pollen [43], S. cerevisiae [30,31], A. nidulans [32] and D. melanogaster [53] cleave nucleic acids endo^exonucleolytically with the formation of oligonucleotides and a small amount of mononucleotides, whereas the nucleases from A. espejiana [12] and B. subtilis [25] degrade nucleic acids in an exo^endo fashion with mononucleotides as the main product of hydrolysis. BAL 31 nuclease hydrolyzes ssDNA endonucleolytically and shortens the linear duplex DNA from both 3P- and 5P-ends. However, the enzyme isolated from N. crassa mitochondria showed distributive endonuclease activity towards ssDNA but processive exonuclease activity towards dsDNA [24]. In addition, an endo^exonuclease from S. cerevisiae, in£uenced by the RAD52 gene, showed a non-processive endo mode of action on ssDNA and weakly processive exonucleolytic activity on dsDNA resulting in the formation of oligonucleotides of varying chain length [30]. In contrast, A. nidulans nuclease showed endo and exo modes of action on both ss and dsDNA [32]. Tobacco pollen nuclease, apart from exhibiting an endo mode of action on ssDNA and exo mode with dsDNA, also showed endonucleolytic activity on dsDNA leading to the formation of a typical 58-bp long fragment [43]. The exo mode of action of N. crassa endo^exonuclease [24] is di¡erent from those exhibited by typical exonucleases like snake venom phosphodiesterase [136], spleen phosphodies-

FEMSRE 729 5-12-01

596

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

terase [137] and A. sydowii nuclease [138], in that the hydrolytic products of N. crassa enzyme consist of di-, triand tetranucleotides. The end-products of hydrolysis of DNA and RNA, by sugar non-speci¢c endonucleases, are oligonucleotides and/or mononucleotides with 5P- or 3P-phosphoryl termini. With the exception of S. aureus [7] and B. subtilis nucleases [25], which produce 3P-mononucleotides, all the other nucleases produce 5P-mononucleotides and/or oligonucleotides with 5P-PO4 and 3P-OH termini. However, wheat chloroplast nuclease hydrolyzes ssDNA endonucleolytically, liberating oligonucleotides with 3P-OH and 5P-PO4 termini while oligonucleotides liberated from RNA hydrolysis have 3P-PO4 and 5P-OH termini [46]. Rye germ ribosomal nuclease I, on the other hand, liberates oligonucleotides ending in 3P-OH and 5P-PO4 from RNA and 3P-PO4 and 5P-OH from ssDNA and dsDNA [47].

ated with both F and S species against 5P-terminated single-stranded tails generated by the exonuclease action [82,145]. The exonuclease activity of the S form, on duplex DNA, decreased with increasing G+C content, whereas the action of F form was not very dependent on the base composition [146]. BAL 31 nuclease F species degraded linear duplex RNA in a terminally directed manner, resulting in the removal of nucleotides from both 3Pand 5P-ends. In comparison, the S species showed very little activity against duplex RNA [147]. Subsequently, Lu and Gray [148] showed that both forms degrade ssDNA in a processive manner from the 5P-end as against the 3P-termini for the duplex DNA. Endo^exonucleases from B. subtilis [149], N. crassa mitochondria [24], S. cerevisiae [31], A. nidulans [32], and D. melanogaster [53] also showed 5PC3P-directed exonuclease action on duplex DNA.

10. Mechanism of action

11. Action on polynucleotides

Mechanisms of hydrolysis of dsDNA by endonucleases have been studied by light-scattering, viscometry and sedimentation analysis [139^141]. In the `double-hit' mechanism, nicks are made at random sites in each strand at points away from each other and the complete fragmentation of duplex DNA does not occur until the two nicks are opposite to each other (multiple encounter). However, in the `single-hit' mechanism, the nicks on either strand are made at points close or opposite to each other, resulting in the complete scission of the duplex DNA in a single encounter [142]. Melgar and Goldthwait [143] studied the e¡ect of metal ions on the hydrolysis of DNA by pancreatic DNase I and demonstrated that, in the presence of Mg2‡ , the DNA is cleaved predominantly by a double-hit mechanism, whereas with Mn2‡ , Ca2‡ or Co2‡ the mechanism of hydrolysis shifts to single-hit mode. On the contrary, metal ions had no e¡ect on the mechanism of action of nuclease Rsn and it cleaved dsDNA through a single-hit mechanism in the presence of Mg2‡ , Mn2‡ and Co2‡ [40]. In the case of nucleases from yeast mitochondria [28] and S. marcescens [144], it has been suggested that the cleavage of the phosphodiester linkage of native DNA occurs at the same or nearby sites, suggesting a single-hit mechanism. In general, sugar non-speci¢c endonucleases show exoand endonucleolytic activities on both DNA and RNA (Table 3). BAL 31 nuclease (both S and F forms) degrades ssDNA exonucleolytically, whereas dsDNA degradation occurs in a terminally directed manner, in which the removal of nucleotides takes place from both the ends of dsDNA. The ratio of the turnover number for the exonuclease activity of the F form, to shorten the duplex DNA, is approximately 27 þ 5 times higher than that of the S form. Apart from terminally directed exonuclease activity, some endonuclease activity was also found to be associ-

Action of sugar non-speci¢c endonucleases on synthetic polynucleotides revealed that the rate of hydrolysis varies with the source of the enzyme. Wheat chloroplast nuclease hydrolyzed various synthetic polynucleotides in the order of poly(A) s poly(U) s poly(C) and poly(dA) s poly(dT) s poly(dC) but poly(G) and poly(dG) were resistant to cleavage [46]. Staphylococcal nuclease also exhibited a similar preference for the homopolyribonucleotides to that of wheat chloroplast nuclease [105]. However, nuclease I from rye germ ribosomes showed high speci¢city for poly(C), while other ribopolynucleotides were hydrolyzed in the order of poly(A) v poly(U) = poly(G). Although rye germ ribosomal nuclease I hydrolyzed the double-stranded deoxyriboheteropolymer poly(dT).poly(rA) at a very slow rate, it failed to hydrolyze the riboheteropolymer poly(A).poly(U), suggesting its preference for single-stranded nucleic acids [47]. Similarly, barley nuclease hydrolyzed the polynucleotides in the order of poly(C) s poly(U)s poly(A) s poly(A).poly(U) s poly(G) = poly(G).poly(C) [102]. In contrast, rye germ ribosomal nuclease II hydrolyzed the double-stranded polymer poly(I).poly(C) at a higher rate than poly(A).poly(U) followed by single-stranded polymers in the order of poly(U) s poly(A) s poly(C) s poly(I). However, poly(G) and poly(dI).poly(dC) were resistant to cleavage [48]. S. cerevisiae nuclease could degrade poly(dT), poly(U), poly(A) and poly(C) but not poly(G), poly(A).poly(U), poly(AU) and poly-(A).poly(dT) [31]. Endonuclease from potato tubers showed preference for polynucleotides in the order of poly(U) s poly(A) s poly(I) s poly(C), whereas the enzyme from rat liver nuclei [50] hydrolyzed these substrates in the order of poly(U) s poly(C) s poly(A). A. agilis nuclease hydrolyzed only poly(A) whereas poly(U), poly(C) and poly(G) were resistant to cleavage. While a 10-fold and 30-fold excess enzyme was required for signi¢cant hydro-

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

lysis of poly(U) and poly(C), respectively, poly(G) was resistant to cleavage even at higher enzyme concentrations [21]. Similarly, nuclease Rsn hydrolyzed only poly(A) while poly(U), poly(C) and poly(G) were resistant to cleavage. However, detectable cleavage of poly(U) and poly(C) could be observed only on prolonged incubation with excess enzyme [106]. S. marcescens kums 3958 nucleases hydrolyzed poly(A) slowly but poly(I).poly(C) was degraded at a higher rate followed by poly(A).poly(U) and poly(dG).poly(dC). However, poly(G), poly(C), poly(U), poly(dT), poly(dA) and poly(dA).poly(dT) were resistant to hydrolysis [23]. On the contrary, Serratia nuclease cleaved the duplex form in the order of poly(dG). poly(dC) s poly(A).poly(U) s poly(dI).poly(dC) s poly(I). poly(C) but poly(dA).poly(dT) was resistant to hydrolysis [126]. Similarly, Anabaena nuclease could readily hydrolyze poly(dG).poly(dC) but poly(dA).poly(dT) was resistant to cleavage, suggesting the preference of the enzyme for the former [118]. 12. Action on plasmid DNA Cleavage of closed circular DNA is considered as one of the main characteristics of endonucleases. In plasmid and phage, closed circular duplex DNA exists in a supercoiled form as a consequence of torsional strain, which at su¤ciently high negative superhelical density promotes unwinding of helical twists [150]. Non-speci¢c endonucleases nick supercoiled DNA (Form I) to give rise to linear duplex DNA (Form III) via the formation of nicked circular DNA (Form II). Thus, N. crassa (mitochondrial and vacuolar) nucleases could readily convert Form I DNA to Form II DNA but its subsequent conversion to linear duplex DNA (Form III) occurred at a slow rate [24]. Similar observations were made with nucleases from A. espejiana [12], barley aleurone layers [102], S. cerevisiae [30], A. nidulans [32], Leishmania sp. [37] and S. antibioticus [39]. In the case of N. crassa mitochondrial nuclease, the enzyme action can be controlled by adjusting the concentrations of Mg2‡ in the reaction mixture. At low enzyme concentrations, in the presence of 0.1 mM Mg2‡ , it exhibits strict endonuclease activity and high speci¢city for Form I DNA. However a 4^8-fold excess enzyme, in the presence of 10 mM Mg2‡ , accelerated the conversion of Form II DNA to Form III DNA followed by exonucleolytic degradation of Form III DNA [24]. D. melanogaster endonuclease converted Form I DNA rapidly to Form III DNA with very little accumulation of Form II DNA. The appearance of low amounts of Form II DNA was attributed to the preferential nicking on one strand before it is linearized [53]. Similarly, low concentrations of nuclease Rsn converted Form I DNA to Form III DNA via Form II DNA. However, with increase in the incubation time, degradation of Form III DNA was observed. Additionally, the conversion of Form I DNA to Form III

597

DNA and its further degradation in the presence of Co2‡ was slow in comparison to Mg2‡ and Mn2‡ , suggesting the relatively low preference of the enzyme for dsDNA in the presence of Co2‡ [40]. Leishmania nuclease, at low enzyme concentrations, generates single base nicks that can be ligated by T4 DNA ligase to yield covalently closed circular DNA [37]. Similarly, S. pombe endonuclease also produces nicks which are resealable by T4 DNA ligase [35]. In contrast, the nicks generated in supercoiled DNA by BAL 31 nuclease (F and S forms) could not be ligated back to covalently closed DNA, since they were extended into gaps by the exonuclease action of these enzymes. However, the ligation of the gaps could be achieved by carrying out the reaction using T4 DNA ligase in combination with T4 DNA polymerase [151]. BAL 31 nuclease exhibited enhanced endonucleolytic activity on netropsin-bound supercoiled plasmid DNA with no detectable exonucleolytic activity [152]. It is known that intercalating agents change the superhelical density of plasmid DNA in the order of less negatively supercoiledCrelaxedCpositively supercoiled. Moreover, negatively supercoiled DNA is known to form stably unwound DNA conformations including ZDNA, cruciform and homopurine^homopyrimidine structures. BAL 31 nuclease cleaves very highly supercoiled DNA prepared from covalently closed relaxed DNA (Form I³) with EtBr. Initial nicking rates of PM2 Form I DNA by BAL 31 nuclease are readily measurable at superhelical densities as low as 30.02 and the nicking activity on positively supercoiled DNA becomes detectable at superhelical densities between 0.15 and 0.19 [81]. It was also noted that non-stressed (i.e. non-supercoiled), nonmodi¢ed duplex DNAs are resistant to endonucleolytic action [153]. 13. Cleavage preference Staphylococcal nuclease hydrolyzed denatured DNA at a faster rate than native DNA, suggesting that the rate of cleavage depends on the conformation of the substrate [154]. Based on the ¢ndings of Felsenfeld and Sandeen [155] that dA^dT-rich regions of native DNA undergo denaturation at somewhat lower temperatures (i.e. below the average Tm ) than regions containing higher proportions of dG^dC nucleotide pairs, von Hippel and Felsenfeld [108] postulated that at low temperatures dA^dT regions undergo local opening and closing or strand separation reaction (`breathing') to a greater extent than dG^dC-rich regions of DNA. This `breathing' phenomenon facilitates the formation of functional enzyme^substrate complexes and accounts for the relative predominance of deoxyadenylic and thymidylic acid residues in the early digestion products of DNA by staphylococcal nuclease [108]. Additionally, Cuatrecasas et al. [105] ob-

FEMSRE 729 5-12-01

598

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

served that the 5P-mononucleotides dA and dT are strong inhibitors of staphylococcal nuclease and suggested it to be the basis of strong binding of the enzyme to dA^dTrich regions, thereby accounting for the release of deoxyadenylic and thymidylic acid residues in the early part of hydrolysis. This led Wingert and von Hippel [109] to suggest that the speci¢city of the enzymatic attack, on native DNA, depends on the conformational motility (`breathing' or `dynamic' structure) of DNA molecule and that denatured DNA is preferentially attacked when both denatured and native DNA are present. This is supported by the observation that staphylococcal nuclease binds to an exposed single-stranded region and nicks transiently melted base pairs in duplex DNA [15]. Moreover, the formation of single-stranded regions during the course of digestion of native DNA was correlated to ss-cut-triggered local unwinding of the double helix. The above phenomenon has been observed in DNA degrading enzymes capable of producing single-strand nicks [156]. The di¡erent pathways for the formation of ssDNA regions are depicted in Fig. 4. The occurrence of gaps in plasmid DNA by the action of BAL 31 nuclease also supports the formation of singlestrand regions in native DNA by DNA degrading enzymes. Similar observations were also made with N. crassa endo^exonuclease [24]. Meiss et al. [144] studied the cleavage of PCR-generated oligonucleotides by Serratia nuclease and demonstrated that the enzyme cleaves preferentially the GC-rich regions (particularly d(G).d(C) tracts) in dsDNA and avoids d(A).d(T) tracts. It was also shown that, compared to DNase I, Serratia nuclease is more non-speci¢c and at-

tacks a particular substrate more evenly under standard reaction conditions. Though the non-speci¢c nature of cleavage is maintained in the presence of high ionic strength or DMSO, addition of urea made the enzyme more selective and this led the authors to suggest that Serratia nuclease could be sensitive to global features like the width of the minor groove [144]. Additionally, experiments with synthetic model substrates (oligonucleotides) showed that Serratia nuclease prefers purine-rich oligonucleotides, probably because it adopts a helical structure with pronounced base stacking. However, cleavage speci¢cities with homo- and heteroduplexes indicated that the A-form of nucleic acid is preferred over the Bform [157]. Similarly, endonuclease from Anabaena sp. also preferred A-form DNA [118]. In the case of endonuclease G from B. taurus, it was shown that the enzyme preferentially cleaves (dG)n .(dC)n tracts in DNA [51,77]. Synthetic polynucleotides, namely poly(dC)n .poly(dG)n and the oligomer d(GGGGCCCC), adopt a right-handed structure similar to classical A-form DNA [158,159]. The susceptibility of these oligonucleotides to endonuclease G led Ruiz-Carrillo and Renaud [51] to conclude that the enzyme recognizes A-type DNA. Like Serratia [157] and Anabaena [118] nucleases, endonuclease G does not cleave d(A).d(T) and (dG.dA)34 .(dT.dC)34 . However, endonuclease G hydrolyzes (dG)24 .(dC)24 tracts in supercoiled DNA, in a bimodal way every 9^11 nucleotides, with the maxima in one strand corresponding to minima on the opposite strand, suggesting that the enzyme binds preferentially to one side of the double helix [51]. The segments of (dC^dG) in DNA restriction fragments and in recombinant plasmids adopt a left-handed conformation in high salt solution, while the neighboring regions of natural sequences remain in right-handed helices [160]. Kilpatrick et al. [161] showed that BAL 31 nuclease cleaves the B^Z junction in the presence of high salt concentration but it does not cleave DNA under conditions where (dC^dG)n blocks exist in the B conformation. 14. Base speci¢city

Fig. 4. Mechanisms of generation of ssDNA regions. The possible modes (a^e) of generation of ssDNA regions are depicted. The arrows indicate the cleavage sites of single-strand-speci¢c enzymes (reproduced with permission from Galcheva-Gargova et al. [156]).

The majority of sugar non-speci¢c endonucleases reported so far are base non-speci¢c, but the enzymes (Sm1 and Sm2) from S. marcescens [162] and tea leaves [42] showed some base speci¢city during the initial stages of hydrolysis. Nucleases Sm1 and Sm2 from S. marcescens preferred uracil and guanosine residues respectively in the polynucleotide chain [162]. This di¡erence in the base speci¢city was correlated to the di¡erence in the N-terminal tripeptide fragment [163]. Tea leaf nuclease hydrolyzed RNA with preferential liberation of 5P-AMP and 5PGMP, but 5P-UMP and 5P-CMP were liberated after a considerable lag period. Although hydrolysis of DNA gave similar results, 5P-dCMP was detected only after exhaustive digestion of the substrate [42]. These observations

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

indicated the preference of the enzyme for purine nucleotides. Interestingly, nuclease Rsn did not exhibit any base preference and cleaved both ss and dsDNA in a non-speci¢c manner [88] but it showed high preference for adenylic acid linkages in RNA [106]. Action of barley aleurone layer nuclease on dinucleoside monophosphates revealed a strong preference for purine nucleosides as the 5P-residue followed by uridine as the 3P-residue. Similarly, staphylococcal nuclease hydrolyzed various dinucleoside monophosphates in the order of d-TpT s d-CpC s dGpG. It was observed that the presence of 5P-phosphate in the dinucleotides brought about a 25-fold increase in the activity, compared to 5-fold when the phosphate was present at the 3P-position [164]. 15. Associated phosphomonoesterase activity Among sugar non-speci¢c endonucleases, only those from potato tubers [41], barley aleurone layers [102], tea leaves [42] and rye germ ribosomes [47] exhibit 3P-phosphomonoesterase activity. Interestingly, all the aforementioned enzymes, except rye germ ribosomal nuclease I [47], are 3P-monoribonucleotides-speci¢c. In addition, some enzymes exhibit a base preference for the hydrolysis of various 3P-monoribonucleotides. Thus, tea leaf nucleases (A and B) hydrolyzed various 3P-monoribonucleotides in the order of A s G s U s C [42], whereas potato tuber nuclease hydrolyzed them in the order of A s U s G s C [41]. In contrast, barley aleurone layer nuclease readily hydrolyzed 3P-AMP and 3P-GMP while 3P-CMP and 3P-UMP were resistant to cleavage [102]. 16. Structure and function The amino acid sequences of nine sugar non-speci¢c endonucleases, derived from their gene sequences, showed that most of them are made up of a single polypeptide chain (Fig. 5). Based on the structural and functional similarity, endonucleases from Anabaena sp. [33], S. cerevisiae [165], S. racemosum [34], C. echinulata [38], S. pombe [35] and B. taurus [61] were shown to belong to the Serratia family of non-speci¢c endonucleases. In all the enzymes, except staphylococcal nuclease, the active site residues are conserved (Fig. 5), including the DRGH motif which contains the active site histidine [119,166,167]. Among them, C1 nuclease from C. echinulata showed 44% and 42% sequence homology with S. cerevisiae and S. pombe nucleases, respectively [38]. Staphylococcal nuclease consists of a single polypeptide chain of 149 amino acids with no disul¢de bonds or free cysteine [83]. Shrimp nuclease is also a single polypeptide containing an open reading frame encoding a putative 21residue signal peptide and a 381-residue mature protein. The polypeptide is cross-linked by ¢ve disul¢de bonds

599

[168]. C1 nuclease from C. echinulata is a monomeric protein containing 252 amino acids cross-linked by a single disul¢de bond [38]. Similarly S. racemosum nuclease, a glycoprotein, is made up of a single polypeptide chain of 250 amino acids with a single disul¢de bond and the carbohydrate moiety is attached to Asn134 [169]. Serratia nuclease is produced as a 266-amino acid pre-protein with a signal peptide consisting of ¢rst 21 residues [170,171]. Among the two major isoforms produced by S. marcescens [23,96], with near identical biochemical properties, the 245-amino acid mature nuclease (Sm2), with a Mr of 26.7 kDa, is the result of a typical signal sequence processing. The second isoform (Sm1) lacks the ¢rst three N-terminal amino acids [96,163,172,173]. Although a large number of sugar non-speci¢c endonucleases have been reported to date, structural studies have been limited to staphylococcal nuclease [174^176] and Serratia nuclease [85,166,177,178]. The crystal structure of staphylococcal nuclease, in the presence of Ca2‡ and î resolution, showed the involvement pdTp at 1.5^1.6 A of 30 residues in three separate sections of the helix and, in addition, about 24 residues form a three-stranded section of anti-parallel L-sheet. Residues 44^53 were found to be somewhat disordered and formed a highly solvent-exposed large loop (omega-loop). Moreover, the structure showed the presence of an inhibitor pocket which is predominantly neutral or hydrophobic. In addition, the inhibitor pocket also contained several residues, which speci¢cally participate in binding of calcium ion and the nucleoside diphosphate [174^176]. The crystal structure of Serratia nuclease, a dimeric enzyme, was ¢rst solved î by Miller et al. [177], and it was then re¢ned at at 2.1 A î and 1.1 A î resolution by Lunin et al. [178] and 1.7 A Shlyapnikov et al. [179], respectively. The crystal structure revealed that the dimer consists of two identical monomers and each monomer contains the active site. Moreover, each monomer consists of a central six-stranded anti-parallel L-sheet £anked on one side by a helical domain and on the opposite side by a dominant helix and a very long coiled loop. The crystal structure also showed that the cleft between the long helix and the loop, near His89, may contain the active site [177]. 16.1. Active site Considerable work has been done on the substrate speci¢city and mode of action of sugar non-speci¢c endonucleases but very little information is available regarding the nature of their active site. S. aureus nuclease bound 5P-mononucleotides in a ratio of 1:1 and was completely dependent on the presence of Ca2‡ . Moreover, binding of 5P-mononucleotides and pdTp to the enzyme was a¡ected by low amounts of DNA and RNA, suggesting the presence of a common catalytic site for the hydrolysis of both the substrates [180]. Studies using amino and nitrotyrosyl derivatives of Tyr85 and Tyr113 indicated that they are in

FEMSRE 729 5-12-01

600

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

Fig. 5. Sequence alignment of non-speci¢c endonucleases. Sma ^ S. marcescens, Asp ^ Anabaena sp., Bos mosum, Spo ^ S. pombe, Cun ^ C. echinuclata, Shr ^ shrimp hepatopancreas and Sta ^ S. aureus. The each line. Identical amino acid residues (excluding those of S. aureus) having v50% sequence similarity homology are boxed. The DRGH motif is depicted in bold face. The conserved active site residues are ments were made based on the data from Ho et al. [38], Friedho¡ et al. [166,167] and Wang et al. [168].

FEMSRE 729 5-12-01

^ B. taurus, Sce ^ S. cerevisiae, Sra ^ S. racenumbers on the right give the last residue of are shaded and regions exhibiting substantial marked with an asterisk. The sequence align-

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

601

Fig. 5 (continued).

stereochemical proximity and act in a concerted manner [181]. Through site-directed mutagenesis it was established that Tyr85 has a more direct role in the active site than Tyr113 or Tyr115 [182]. Enzyme crystals, obtained in the presence of Ca2‡ and pdTp, revealed the involvement of Lys84 and Tyr85 in the catalytic activity and that they form hydrogen bonds with the 3P-phosphate of the inhibitor, whereas Arg35 and Arg87 form hydrogen bonds with the 5P-phosphate. Carboxylate ions of Glu43, Asp21 and Asp40 serve as ligands for the binding of Ca2‡ [174]. Cotton et al. [175] suggested that pdTp, in particular its 5Pphosphate, and the activating Ca2‡ are precisely and rigidly locked into the position at the hydrolytic site. It was also proposed that while Glu43 acts as a general base, Arg35 and Arg87 are involved in the neutralization of the charges on the phosphates and Ca2‡ performs the dual role of aiding catalysis and stabilization of the trigonal bipyramid transition state. Crystal structure studies of

î resolution con¢rmed the staphylococcal nuclease at 1.6 A above observations [176]. Based on the sequence homology of related nucleases [20,166], mutational analysis of the conserved amino acid residues [166] and crystal structure [85,177], Friedho¡ et al. [167] proposed that the active site of Serratia nuclease comprises at least four residues, namely: Arg57, His89, Asn119 and Glu127. Since the mutants of active site residues showed a similar e¡ect on the hydrolysis of both DNA and RNA, the authors suggested the existence of a common catalytic site for the hydrolysis of both substrates. Furthermore, it was observed that among all mutants of His89, only H89N showed measurable enzyme activity in addition to a decrease in the kcat , suggesting the probable role of His89 in catalysis. However, the similar substrate binding e¤ciency of the wild-type enzyme and mutant H89A, of the non-cleavable modi¢ed substrate (a decamer in which phosphates are replaced by

FEMSRE 729 5-12-01

602

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

phosphorothioates), suggested that His89 is not involved in substrate binding. It was also demonstrated that His89 has to be deprotonated to function as a general base in the catalytic activity [167]. The close proximity of Arg57 and Arg87 to the phosphate backbone in the substrate [177], coupled with the decrease in the kcat of the Arg57 mutants and with no change in the Km , suggested the involvement of Arg57 in catalysis. Moreover, the signi¢cant decrease in the kcat of the Arg57 mutant was correlated to its involvement in positioning and polarizing the phosphate of the scissile phosphodiester bond and/or stabilization of the transition state. Asn119 was implicated in catalysis since mutants of Asn119 showed a decrease in their kcat and not in the Km [166,167]. Subsequently Miller et al. [183] established the involvement of Asn119 in metal binding. Compared to the native enzyme, the alanine mutants of Glu127 showed reduced activity while the mutants E127D and E127Q showed similar activities to that of the native enzyme, indicating that Glu127 might be participating indirectly in the hydrolysis of the substrates [167]. Non-speci¢c endonuclease from Anabaena sp. showed nearly 30% sequence homology with Serratia nuclease and this was restricted mainly to the active site regions and the central six-stranded L-sheet of the enzyme. Based on the three dimensional structure of S. marcescens nuclease and mutational analysis of the catalytically active residues, a structural model for the Anabaena nuclease was proposed. Accordingly, it was suggested that His124, Asn155 and Glu163 (corresponding to His89, Asn119 and Glu127 of S. marcescens nuclease) are involved in the catalytic activity of the enzyme. Additionally, through mutational analysis it was demonstrated that Asn155 and Asp121 are involved in the coordination of the metal ion cofactor. Subsequently cleavage studies with deoxythymidine 3P,5Pbis-(p-nitrophenyl phosphate) revealed the role of His124 as a general base and that of Glu163 in assisting the general acid [119]. Chemical modi¢cation studies on nuclease Rsn, from R. stolonifer, showed the involvement of two histidine, one tryptophan and two carboxylate residues in the catalytic activity of the enzyme. Substrates of nuclease Rsn, viz. DNA and RNA, could not protect the enzyme against DEP- and EDAC-mediated inactivation, whereas substrate protection was observed in the case of NBS-mediated inactivation of the enzyme. Km and kcat values of the partially inactivated enzyme samples suggested that while histidine and carboxylate are involved in catalysis, tryptophan is involved in substrate binding. Furthermore, £uorescence quenching studies on native and modi¢ed nuclease Rsn, using metal ions, indicated the involvement of carboxylate in metal binding [184]. Based on the sequence similarity with S. marcescens and site-directed mutagenesis, the involvement of His87, His85 and His211 in the catalytic activity of endonucleases from C. echinulata [38], S. racemosum [169] and shrimp hepatopancreas [168], respectively, was demonstrated. Moreover,

the probable involvement of Asn241 in catalysis and Lys210 and Arg253 in substrate binding was postulated for shrimp hepatopancreas nuclease [168]. In the case of the mitogenic factor secreted by S. pyogenes, His122 was shown to be the residue essential for the nuclease activity [36]. 16.2. Coordination and function of metal ions Staphylococcal nuclease is a Ca2‡ -dependent non-specific enzyme, while S. marcescens nuclease requires Mg2‡ for its activity. Extensive studies on these enzymes have provided insights into the substrate binding, metal interaction and catalytic mechanism. In case of staphylococcal nuclease, Ca2‡ is coordinated directly to Asp21 and Asp40, while Glu43 is coordinated to the calcium ion through a water molecule. Additionally, the calcium ion was found to coordinate with two additional water molecules (Fig. 6) [174,176,185]. The roles of Asp21 and Asp40 were further delineated by electron-spin paramagnetic resonance studies, in which Asp21 was found to in£uence the binding of Ca2‡ more strongly than Asp40 [186]. Through the crystal î resolution of the D21E ternary comstructure at 1.95 A plex, where D21E is bound to both Ca2‡ and pdTp (transition state analogue), it was demonstrated that Ca2‡ in the active site binds to Glu21 via bidentate coordination and to Glu43 through an inner sphere coordination. The cooperativity of the metal ion and the inhibitor (pdTp) in the ternary complex was also demonstrated [187]. On the other hand, in Serratia nuclease the catalytic magnesium binding site is located between the long helix (residues 116^135) and the loop extending from residues

Fig. 6. Schematic view of the binding site of staphylococcal nuclease in the enzyme^pdTp^Ca2‡ complex. The heavy dots represent regions of electron density assigned as water molecules. Hydrogen bond and ionic interactions are shown as dotted lines. The interaction between Ca2‡ and Val41 is through the carbonyl oxygen of the peptide bond (reproduced with permission from Cotton et al. [175]).

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

Fig. 7. Schematic view of the magnesium binding site of Serratia nuclease (reproduced with permission from Miller et al. [183]).

50 to 114, which contains the catalytically essential His89. It was shown that Mg2‡ bound to Asn119 (the only protein ligand) is associated with ¢ve water molecules to complete an octahedral coordination complex (Fig. 7). Glu127 and His89 are located nearby and each is hydrogenbonded to water molecules in the coordination sphere. Kinetic and site-speci¢c mutagenesis studies suggested that this metal^water cluster contained the catalytic metal ion. The residues involved in the formation of metal^water cluster coordination are conserved in enzymes homologous to Serratia nuclease. Additionally, it was demonstrated that Serratia nuclease is a unique enzyme of the endonuclease family in which the amide side-chain acts as the sole protein ligand for magnesium coordination [183]. Though Lunin et al. [178] proposed that Asp86 is involved in Mg2‡ coordination, Miller et al. [183] demonstrated that Asp86 interacts indirectly with the metal binding site by forming hydrogen bonds with the amide nitrogen of Asn119 and Gln120. Similarly, Meiss et al. [119] showed that the metal cofactor is bound to Asp121 in addition to Asn155, which is the primary ligand for binding the cofactor. 16.3. Enzyme^substrate interactions In staphylococcal nuclease, binding of either Ca2‡ or the substrate/inhibitor to the enzyme is not mutually exclusive [180,188]. Cuatrecasas et al. [180] showed that Ca2‡ does not bind to the enzyme in the absence of nucleotide or substrate and hence suggested that Ca2‡ and nucleotide bind to the enzyme in a unique complex formed only when all three components are present. Before the crystal structures of staphylococcal nuclease were solved, Cuatrecasas et al. [189] postulated that the active site of the enzyme consists of three phosphate binding `subsites'

603

(namely P1, P2 and P3) and the binding of oligonucleotides occurs predominantly by ionic interactions. The contribution of subsites in the binding of substrates/inhibitors to form a complex was shown to be in the order of P1 s P2 s P3. Also, the cleavage speci¢city of the hydrolytic site, visualized as an integral part of the P1 subsite, was shown to be in£uenced by the presence of nucleoside on the 5P-side rather than the 3P-side [189]. Weber et al. [190] postulated that the binary enzyme^dTdA complex is one of the intermediates leading to the active ternary enzyme^metal^substrate complex. Based on the conformational changes between the binary and ternary complexes it was demonstrated that the metal-induced changes, both at the attacked phosphorus and at the leaving group of the enzyme-bound substrate, may contribute to catalysis. Stanczyk and Bolton [191], using crystal structures of the ternary complexes of staphylococcal nuclease with Ca2‡ and pdTp, showed that the 5P-phosphate of pdTp interacts with the solvent-inaccessible Arg35 and Arg87, whereas the 3P-phosphate and Ca2‡ interact with the solvent-accessible O-ammonium group of Lys84 and the phenolic hydroxyl group of Tyr85. It was also observed that the interaction of a terminal 3P-phosphate for exonucleolytic activity di¡ers slightly from the interaction with an internal phosphodiester linkage required for the endonucleolytic cleavage. Subsequently, using the ternary complexes of the enzyme with pdTp, pdGp and p-nitrophenyl-pdTp, the above authors [191] demonstrated that the conformational features of the bound nucleic acid determine the di¡erences in the observed catalysis rather than the nucleotide itself. Similar observations were also reported by Grissom and Markley [192]. In the case of Serratia nuclease, the minimum substrate recognized is a pentanucleotide containing phosphate at the 5P-position [193]. Friedho¡ et al. [133] demonstrated that the 5P-phosphorylated pentamers are cleaved predominantly at the second phosphodiester bond and to a lesser extent (5%) at the third phosphodiester position. Additionally, at high enzyme concentrations, cleavage of the tetramer was observed at the ¢rst and second phosphodiester positions with no detectable cleavage at the third phosphodiester bond. Based on the above observations, the authors concluded that a phosphate 3P to the scissile phosphodiester bond is important for the cleavage. This conclusion is supported by the cleavage of the arti¢cial substrate deoxythymidine-3P,5P-bis-(p-nitrophenyl phosphate) by S. marcescens nuclease [134]. Furthermore, Serratia nuclease is not active against methylphosphonate-substituted pentanucleotides suggesting that the negative charges and/or the hydrogen bond acceptors on the phosphate to be attacked, as well as on the neighboring phosphates, are required for binding and correct orientation of the substrate in the active site of the enzyme [133,194]. The requirement of phosphate groups for the e¡ective binding of the substrate to the active site of the enzyme was found to be similar for both S. aureus and S. marcescens nucle-

FEMSRE 729 5-12-01

604

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

ases. Moreover, cleavage experiments with polydeoxyadenylates, containing a single phosphorothioate at di¡erent positions, indicated that Serratia nuclease hydrolyzed the Rp -diastereomer at a higher rate than the Sp -diastereomer. Hence, the authors opined that the Sp orientation prevents cleavage at the phosphorothioate group and retards the cleavage at a position 5P to the substitution, whereas the Rp -phosphorothioate group enhances the cleavage at the 5P-position adjacent to the site of substitution [133]. Interestingly, the role of Trp123 in substrate binding was established by kinetic studies involving mutant and wild-type enzymes [157]. Similarly, in the case of the Anabaena nuclease Meiss et al. [119] demonstrated the involvement of Trp159 in substrate binding. Moreover, studies on the mechanism of inhibition of Anabaena nuclease (NucA) by its inhibitor (NuiA), using several active site mutants of NucA, suggested that Glu163 of NucA might be participating in cofactor binding or protonation of the leaving group as the target amino acid involved in the inhibition of the enzyme. Based on these observations, it was concluded that Glu163 of the enzyme interacts with the positively charged amino acids of the inhibitor, while Arg93, Arg122 and Arg167 of the enzyme interact with the negatively charged amino acids of the inhibitor [119]. 16.4. Water-assisted metal ion catalysis The crystal structures of staphylococcal nuclease [174^ 176] and Serratia nuclease [178,183] revealed the presence of water molecules adjacent to the metal ions. In both cases, the reaction is triggered by the water molecule which acts as the nucleophile and attacks the scissile phosphodiester bond. The reaction mechanism, for staphylococcal nuclease, based on the crystal structure [175,176], kinetic studies [195,196] and site-directed mutagenesis [186,197^201], is depicted in Fig. 8. The reaction mechanism involves the Ca2‡ activator bound in a septacoordinate complex [176] to Asp21, Asp40 and the amide carbonyl group of Thr41 as well as the 5P-phosphate of the competitive inhibitor (pdTp), with the remaining three ligands probably being the water molecules. The attacking water molecule is presumed to be near the Ca2‡ , either in direct coordination or in the second sphere of Glu43, to permit its carboxylate group to function as a general base [199,200]. Upon nucleophilic attack, the phosphodiester linkage of the substrate is converted into a trigonal^bipyramid intermediate stabilized by Arg87 [186,201], which also acts as the general acid to protonate the 5P-oxygen of the leaving nucleoside. Carboxylate groups are known to be e¡ective nucleophilic catalysts in the intramolecular hydrolysis of phosphate esters [202]. The mechanism of staphylococcal nuclease suggested by Cotton et al. [175] requires that the Qcarboxylate group of Glu43 acts as a general base, facilitating the attack of water on the scissile phosphodiester bond. Judice et al. [203], while probing the mechanism of

Fig. 8. Reaction mechanism of staphylococcal nuclease (reprinted with permission from Weber et al. [190], Copyright 1991, Americal Chemical Society).

staphylococcal nuclease with unnatural amino acids, suggested that Glu43 plays a more complex structural role in catalysis rather than acting as a general base. To ascertain the role of Glu43 as a general base, Loll et al. [204] solved the crystal structure of the ternary complex of staphylococcal nuclease in the presence of Co2‡ , as the structure is very similar to the structure obtained with Ca2‡ with little or no distortions of the groups involved in the catalytic activity. It was observed that Co2‡ occupies the site identical to the Ca2‡ binding site and both the metals show a similar interaction with the inner sphere of Glu43, indicating that Glu43 is still fully capable of activating the water molecule to act as a general base. Additionally, it was noted that the inner sphere interaction of Ca2‡ with Asp21 was missing in the Co2‡ structure, thus establishing the fact that the loss of catalytic activity of Co2‡ -substituted nuclease is probably due to its inability to bind the inner sphere water [204]. As mentioned earlier, Glu43 in staphylococcal nuclease is situated at the base of the solvent-exposed and conformationally mobile omega-loop in the active site [174]. High resolution X-ray structures revealed that the substitution of Glu43 by aspartic acid signi¢cantly changes the structure of the omega-loop and reduces the interaction of Ca2‡ with its active site ligands, resulting in a diminished hydrogen-bonded network of the water molecules [205]. Mehdi and Gerlt [206] showed that the hydrolysis of substrates by staphylococcal nuclease proceeds with the inversion of con¢guration at phosphorus, aided by Ca2‡ , which assists in the proper positioning of the carboxylate group via an intervening water molecule and neutralizes the phosphate ester charge by a direct ionic interaction. Similar observations were made by Potter et al. [207,208] with the S1 and P1 nucleases. The rate determining step for the hydrolysis of DNA/RNA by staphylococcal nuclease at pH 9.5 was shown to be substrate binding and product dissociation rather than cleavage of the phosphodiester bond. However, under similar

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

conditions with the E43D mutant, in which the putative active site general base catalyst Glu43 is replaced by aspartic acid, cleavage of the phosphodiester bond becomes the rate determining step [209]. As mentioned earlier, Serratia nuclease also catalyzes the hydrolysis of phosphodiester bonds through a metal ion-mediated mechanism with a water molecule acting as the nucleophile. Using synthetic substrates [134], site-directed mutagenesis [166,167] and crystal structures [177,183], Friedho¡ et al. [210] proposed a `general base model' for the reaction mechanism of Serratia nuclease (Fig. 9A). A similar reaction mechanism has also been postulated by Meiss et al. [119] for the endonuclease from Anabaena sp. (Fig. 9B). In the `general base mechanism' postulated for Serratia nuclease (Fig. 9A), His89 acts as a general base by abstracting a proton from the water molecule thereby activating it for a nucleophilic attack on the phosphorous atom adjacent to the scissile bond, followed by cleavage of the 3P O^P bond [134,177]. Magnesium ion acts as the Lewis acid to stabilize the negative charge on the penta-

Fig. 9. Reaction mechanism of Serratia (A) and Anabaena (B) nucleases (reproduced with permission from Meiss et al. [119]).

605

coordinate phosphate transition state and the leaving group. It has been postulated that Asn119 is probably involved in the binding and correct positioning of the metal cofactor. Arg57 has been shown to be involved in the transition state stabilization [210]. Interestingly, homing endonuclease I-PpoI from Physarum polycephalum [210,211], colicin E9 endonuclease [212] and junction resolvase endonuclease (Endo VII) from phage T4 cells [213] shows an identical active site and has been postulated to have a similar mechanism. Anabaena nuclease follows a similar reaction mechanism, where His124 acts as the general base, Asn155 in binding and correct positioning of the cofactor, and Glu163 in protonation of the leaving group (Fig. 9B). In contrast to Arg57 in Serratia nuclease, the transition state stabilization in Anabaena nuclease is effected by Asp95, either by hydrogen bonding or by binding to a second metal ion [119]. Active site characterization of nuclease Rsn from R. stolonifer showed the involvement of similar residues to those of S. marcescens nuclease in the catalytic activity of the enzyme. Hence the authors suggested that, like S. marcescens nuclease, nuclease Rsn may also follow the metal^water cluster-mediated mechanism for the hydrolysis of DNA and RNA [184]. Miller et al. [183] postulated two schemes for the hydrolysis of phosphodiester bonds by Serratia nuclease, where an unligated water molecule may be directly activated by His89, or the magnesium-bound water is activated by His89. In this the metal ion may alter the pKa of the bound water to produce a more nucleophilic hydroxide ligand, and this activated water molecule may mediate the cleavage of the phosphodiester bond. In the former, the cluster polarizes the phosphate atom, stabilizes the phosphorane and helps protonate the leaving group, whereas in the latter, the cluster polarizes the phosphate atom, stabilizes the phosphorane and helps activate the attacking hydroxide. Although Lunin et al. [178] initially proposed a general acid model for Serratia nuclease, subsequently based on the observation of Kolmes et al. [134] that the E127A mutant, and not H89A, cleaves deoxythymidine 3P,5P-bis(p-nitrophenyl) phosphate (as the protonation of the leaving group is not necessary), Shlyapnikov et al. [179] proposed a consensus mechanism similar to the one proposed by Friedho¡ et al. [210]. In the case of Serratia nuclease, the rate determining step for the hydrolysis of oligonucleotides is substrate binding and/or phosphodiester bond cleavage but not product dissociation [133]. The reaction mechanisms of staphylococcal and Serratia nucleases suggest that the hydrolysis of the phosphodiester bond is very much dependent on the proper orientation of the residues so as to initiate an in-line attack of the water molecule on the scissile phosphodiester bond. Also, the proper orientation is achieved by the concerted action of Asn21, Asn40 and Ca2‡ in the case of staphylococcal nuclease [175,176] and Asn119 and Mg2‡ in Serratia nuclease [177,183]. It is interesting to note that though both en-

FEMSRE 729 5-12-01

606

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

zymes could e¤ciently perform the dual roles of proper orientation and transition state stabilization of the intermediate or the leaving group through a single metal atom, P1 nuclease from P. citrinum, a zinc metalloprotein, required three metal ions for performing a similar role [214]. Accordingly, the scissile phosphate of the substrate binds close to ZnII and the base 5P to the bond to be cleaved stacks against phenylalanine (Phe61) and forms hydrogen bonds with aspartic acid (Asp63). ZnI and ZnIII bridge the water molecule, which acts as the nucleophile attacking the phosphate in-line with the P^O3P bond [215]. The carboxylate group of aspartic acid (Asp45), which also serves as a ligand of ZnI, helps to orient the hydroxide properly for the attack. An arginine residue (Arg48) stabilizes the resulting pentacoordinate transition state and the attacking hydroxide ion along with the leaving oxyanion (O3P) occupy apical positions in this transition state. ZnII plays a crucial role in activating the phosphate and stabilizing the leaving O3P oxyanion. Thus, all the three zinc ions are important for catalysis [214]. 17. Biological role As mentioned earlier, nucleases play an important role in replication, recombination, restriction and repair. Extracellular enzymes have been implicated in scavenging of nucleotides and phosphates for the growth and metabolism. Nucleases have been shown to have a role in the formation of nicks during meiotic recombination. The reduced amounts of endo^exonuclease in rad52 mutants of S. cerevisiae were correlated to its probable involvement in repair and recombination in both mitotic and meiotic cells [30]. The ability of N. crassa endo^exonuclease to act on supercoiled DNA and its ability to initiate single-strand and double-strand breaks in DNA along with the processive exonucleolytic action on circular DNA in the presence of ssDNA binding proteins, led Chow and Fraser [24] to suggest that it might be involved in the production of stable lengths of ssDNA needed for exchanges during recombination and recombinational repair of mitochondrial DNA. In the case of N. crassa the repair de¢cient and UVsensitive mutants, uvs-2, uvs-3, uvs-6 and nuh-4, could not secrete endo^exonucleases. These mutants had a higher level of endo^exonuclease precursor than the wild-type, indicating that they may have some defect either in the protease(s) that control the nuclease level or in the regulation of protease(s). The above mutants were also sensitive to various mutagens and mitomycin C and exhibited a high frequency of spontaneous, recessive lethal mutations and deletions, indicating the involvement of N. crassa nuclease in repair [216]. Moreover, it has been suggested that N. crassa mitochondrial endonuclease may have a role in replication [11]. Siwecka et al. [47] postulated the interaction between rye germ nuclease-I and ribosomes to be a part of the regulatory mechanism of cell metabolism. Ow-

ing to the striking correlation between genetic instability and the presence of poly(dG).poly(dC) tracts in DNA (the preferred substrate of endonuclease G), Ruiz-Carrillo and Renaud [51] suggested a probable role for endonuclease G in recombination. Pollen has been used as a promising host for introducing DNA into higher plants [217,218]. The pollen nucleases from P. hybrida [45] and tobacco [43] prefer ssDNA thereby enhancing the chances of the uptake and expression of native DNA. Recently, Nicieza et al. [39] isolated two extracellular nucleases from S. antibioticus, with Mr values of 18 and 34 kDa, which are nutritionally regulated and reach their maximum activity during aerial mycelium formation and sporulation. Their role appeared to be DNA degradation in the substrate mycelium and supply of building blocks for macromolecular biosynthesis in the aerial mycelium, and they acted in concert with the periplasmic nuclease. Of the two extracellular nucleases, the 18-kDa nuclease appeared to be reminiscent of NUC-18, a thymocyte nuclease assumed to have a key role in glucocorticoid-stimulated apoptosis [219,220]. Interestingly, the N-terminal sequence of the 18-kDa protein showed striking similarity to proteins of the cyclophilin family which degrade DNA in a Ca2‡ /Mg2‡ -dependent manner and their role in apoptosis has been reviewed [221,222]. The mitogenic factor of S. pyogenes has been implicated in pathogenesis similar to other pyrogenic exotoxins, but the role of the nuclease activity exhibited by the mitogenic factor is still not clear [36]. DNA/RNA non-speci¢c nucleases from S. aureus and S. epidermidis are found in a variety of clinical and food infections [223^228]. Similarly, nucleases from Vibrio cholerae [229] and S. marcescens [230] have been postulated to play a role during invasion or establishment of an infection. Matousek et al. [231] showed that pollen RNases, owing to their ability to degrade dsRNA, may function as defense proteins against viral infection as components of a degradation complex which participates in the apoptosis of the tapetal cell layer and for nucleoside re-utilization by the developing pollen. The main role of the associated nucleotidase activity of nucleases is to scavenge nucleotides and phosphates for growth and survival of the organism under environmentally stressed conditions. Brown and Ho [102] opined that barley nuclease, in concert with other acid phosphatases, might be involved in the hydrolysis of remnant nucleic acid in the endosperm for providing the valuable nutrients for heterotrophic embryonic growth. A similar role has been suggested for potato tuber nuclease [41]. 18. Applications Since their discovery, sugar non-speci¢c endonucleases have been extensively used as analytical tools for the determination of nucleic acid structure. The N. crassa nucle-

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

ase was employed for the isolation of pure lac operon [232], isolation of tRNA and rRNA gene hybrids [233,234] and detection of sequence heterology [235]. Staphylococcal nuclease has been used to monitor hybridization reactions with DNA [14] and as a probe for drug binding sites on DNA [19]. The ability of S. aureus and N. crassa nucleases to discriminate between uridine and cytidine under de¢ned conditions has been exploited for rapid RNA sequencing [236]. Wheat chloroplast nuclease, due to its preference for pyrimidines in non-base-paired regions, has been used as a tool for the structural analysis of RNA [16]. The ssDNase activity of BAL 31 nuclease was used for probing regions of altered secondary structures in negatively and positively supercoiled closed circular DNA [18]. Additionally, the dsDNase activity of BAL 31 nuclease which progressively shortens the duplex DNA, from both ends, was exploited for ordering restriction endonuclease-generated DNA fragments [17]. Serratia nuclease, under the trade name of `Benzonase', is used to eliminate nucleic acid contamination from puri¢ed recombinant proteins and single cell protein preparations. The ability of Serratia nuclease to degrade DNA has paved way for its utilization as a killer gene for the self-destruction of microorganisms released into the environment in addition to its use as an anti-viral agent [58]. 19. Conclusions The ability of sugar non-speci¢c endonucleases to recognize a wide variety of nucleic acid structures has led to considerable e¡orts to evaluate their role in di¡erent cellular processes as well as application as analytical tools to study nucleic acid structure. The majority of these enzymes share common properties like multiple activity and metal ion requirement. Although these enzymes have been extensively studied with respect to their catalytic properties, very little attention was paid to their structure^ function relationship. The amino acid sequences of typical sugar non-speci¢c endonucleases from S. marcescens, S. cerevisiae, B. taurus, C. echinulata var. echinulata, S. racemosum, S. pombe, Anabaena sp. and shrimp hepatopancreas showed signi¢cant homology in their primary structure. The comparison of primary sequences, conserved sequences and residues involved in the active site of sugar nonspeci¢c endonucleases will advance our understanding on the evolutionary aspect of these enzymes. Until now, the crystal structures of only two enzymes, namely staphylococcal and Serratia nucleases, have been solved. Solving the three dimensional structures of di¡erent sugar nonspeci¢c enzymes will be of great importance as it will add valuable information regarding the structural homology of these enzymes at the tertiary level. Moreover, the majority of the sugar non-speci¢c endonucleases show the requirement of more than one metal ion for their activity. Some enzymes exhibit synergism when metal ions are used

607

in combination. Interestingly, the sugar non-speci¢c endonuclease from R. stolonifer shows a high preference for ssDNA in the presence of Co2‡ . Hence, site-speci¢c mutagenesis in combination with the crystal structures of these enzymes, in the presence and absence of metal ions, will not only yield information regarding the residues involved in the catalytic activity but also the catalytic mechanism of this class of enzymes. In contrast, sugar non-speci¢c endonucleases from plants do not show an obligate requirement for metal ions for their activity. The nature of the active site and the three dimensional structure of these enzymes will be helpful in comparing the mechanisms of action of metal requiring and metal non-requiring endonucleases. Acknowledgements This work was supported by a grant from the Department of Science and Technology, Government of India, to V.S. This is communication No. 6613 from the National Chemical Laboratory, Pune, India. References [1] Araki, T. (1903) Enzymatic decomposition of nucleic acids. Z. Physiol. Chem. 38, 84^92. [2] Iwano¡, L. (1903) Fermentative decomposition of thymo-nucleic acid by fungi. Z. Physiol. Chem. 39, 31^37. [3] Kunitz, M. (1940) Crystalline ribonuclease. J. Gen. Physiol. 24, 15^ 30. [4] Kunitz, M. (1950) Crystalline desoxyribonuclease I. Isolation and general properties. Spectrophotometric method for the measurement of deoxyribonuclease activity. J. Gen. Physiol. 33, 349^355. [5] Laskowski, M. (1959) Enzymes hydrolyzing DNA. Ann. N.Y. Acad. Sci. 81, 776^783. [6] Laskowski Sr., M. (1967) DNases and their use in the studies of primary structure of nucleic acids. Adv. Enzymol. 29, 165^220. [7] Cunningham, L., Catlin, B.W. and Privat de Gar'lhe, M. (1956) A deoxyribonuclease of Micrococcus pyogenes. J. Am. Chem. Soc. 78, 4642^4645. [8] Schmidt, G. (1955) Nucleases and enzymes attacking nucleic acid components. In: The Nucleic Acids, Vol. 1 (Charga¡, E. and Davidson, J.N., Eds.), pp. 555^626. Academic Press, New York. [9] Bernard, E.A. (1969) Ribonucleases. Annu. Rev. Biochem. 38, 677^ 682. [10] Laskowski, Sr. M. (1982) Nucleases: historical perspectives. In: Nucleases (Linn, S. and Roberts, R.J., Eds.), pp. 1^21. Cold Spring Harbor Laboratory Press, New York. [11] Linn, S. and Lehman, I.R. (1966) An endonuclease from mitochondria of Neurospora crassa. J. Biol. Chem. 241, 2694^2699. [12] Gray Jr., H.B., Ostrander, D.A., Hodnett, J.L., Legerski, R.J. and Robberson, D.L. (1975) Extracellular nucleases of Pseudomonas BAL 31. I. Characterization of single-strand-speci¢c deoxyriboendonuclease and double-strand deoxyriboexonuclease activities. Nucleic Acids Res. 2, 1459^1492. [13] Linn, S. (1982) Tabulation of some well-characterized enzymes with deoxyribonuclease activity. In: Nucleases (Linn, S. and Roberts, R.J., Eds.), pp. 341^357. Cold Spring Harbor Laboratory Press, New York. [14] Kacian, D.L. and Spiegelman, S. (1974) Use of micrococcal nuclease

FEMSRE 729 5-12-01

608

[15] [16]

[17]

[18]

[19]

[20]

[21]

[22]

[23]

[24]

[25]

[26]

[27]

[28] [29]

[30]

[31]

[32] [33]

[34]

[35]

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613 to monitor hybridization reactions with DNA. Anal. Biochem. 58, 534^540. Drew, H.R. (1984) Structural speci¢cities of ¢ve commonly used DNA nucleases. J. Mol. Biol. 176, 535^557. Przykorska, A., Kuligowska, E., Gerhart, E., Wagner, H., Szarkowksi, J.W. and Nordstrom, K. (1989) Two plant nucleases as tools for structural analysis of RNA. Biochemistry 28, 1585^1588. Legerski, R.J., Hodnett, J.L. and Gray Jr., H.B. (1978) Extracellular nucleases of Pseudomonas BAL 31 III. Use of the double-strand deoxyriboexonuclease activity as the basis of a convenient method for the mapping of fragments of DNA produced by cleavage with restriction enzymes. Nucleic Acids Res. 5, 1445^1464. Lau, P.P. and Gray Jr., H.B. (1979) Extracellular nucleases of Alteromonas espejiana BAL 31 IV. The single strand-speci¢c deoxyribonendonuclease activity as a probe for regions of altered secondary structure in negatively and positively supercoiled closed circular DNA. Nucleic Acids Res. 6, 331^357. Fox, K.R. and Waring, M.J. (1987) The use of micrococcal nuclease as a probe for drug-binding sites on DNA. Biochim. Biophys. Acta 909, 145^155. Fraser, M.J. and Low, R.L. (1993) Fungal and mitochondrial nucleases. In: Nucleases, 2nd edn. (Linn, S.M., Lloyd, R.S. and Roberts, R.J., Eds.), pp. 171^207. Cold Spring Harbor Laboratory Press, New York. Stevens, A. and Hilmoe, R.J. (1960) Studies on a nuclease from Azotobacter agilis I. Isolation and mode of action. J. Biol. Chem. 235, 3016^3022. Eaves, G.N. and Je¡ries, C.D. (1963) Isolation and properties of an exocellular nuclease of Serratia marcescens. J. Bacteriol. 85, 273^ 278. Yonemura, K., Matsumoto, K. and Maeda, H. (1983) Isolation and characterization of nucleases from a clinical isolate of Serratia marcescens kums 3958. J. Biochem. (Tokyo) 93, 1287^1295. Chow, T.Y.-K. and Fraser, M.J. (1983) Puri¢cation and properties of single strand DNA-binding endo^exonuclease of Neurospora crassa. J. Biol. Chem. 258, 12010^12018. Kanamori, N., Sakabe, K. and Okazaki, R. (1973) Extracellular nucleases of Bacillus subtilis I. Puri¢cation and properties. Biochim. Biophys. Acta 335, 155^172. Maeda, M. and Taga, N. (1976) Extracellular nuclease produced by a marine bacterium. II. Puri¢cation and properties of extracellular nuclease from a marine Vibrio sp.. Can. J. Biochem. 22, 1443^1452. von Tigerstrom, R.G. (1980) Extracellular nucleases of Lysobacter enzymogenes : production of the enzymes and puri¢cation and characterization of an endonuclease. Can. J. Biochem. 26, 1029^1032. Morosoli, R. and Lusena, C.V. (1980) An endonuclease from yeast mitochondrial fractions. Eur. J. Biochem. 110, 431^437. von Tigerstrom, R.G. (1982) Puri¢cation and characteristics of a mitochondrial endonuclease from the yeast Saccharomyces cerevisiae. Biochemistry 21, 6397^6403. Chow, T.Y.-K. and Resnick, M.A. (1987) Puri¢cation and characterization of an endo^exonuclease from Saccharomyces cerevisiae that is in£uenced by the RAD52 gene. J. Biol. Chem. 262, 17659^17667. Dake, E., Hofmann, T.J., McIntire, S., Hudson, A. and Zassenhaus, H.P. (1988) Puri¢cation and properties of the major nuclease from mitochondria of Saccharomyces cerevisiae. J. Biol. Chem. 263, 7691^ 7702. Koa, H., Fraser, M.J. and Kafer, E. (1990) Endo^exonuclease of Aspergillus nidulans. Biochem. Cell Biol. 68, 387^392. Muro-Pastor, A.M., Flores, E., Herrero, A. and Wolk, C.P. (1992) Identi¢cation, genetic analysis and characterization of a sugar-nonspeci¢c nuclease from the cyanobacterium Anabaena sp. PCC 7120. Mol. Microbiol. 6, 3021^3030. Chen, L.-Y., Ho, H.-C., Tsai, Y.-C. and Liao, T.-H. (1993) Deoxyribonuclease of Syncephalastrum racemosum ^ enzymatic properties and molecular structure. Arch. Biochem. Biophys. 303, 51^56. Ikeda, S., Maeda, N., Ohshima, T. and Takata, N. (1996) Identi¢ca-

[36]

[37]

[38]

[39]

[40]

[41]

[42]

[43] [44]

[45]

[46]

[47]

[48]

[49]

[50]

[51]

[52]

[53]

[54]

[55]

tion and characterization of a mitochondrial endonuclease from yeast, Schizosaccharomyces pombe. Biochem. Mol. Biol. Int. 40, 1017^1024. Iwasaki, M., Igarashi, H. and Yutsudo, T. (1997) Mitogenic factor secreted by Streptococcus pyogenes is a heat-stable nuclease requiring His122 for activity. Microbiology 143, 2449^2455. Mittra, B., Sadhukhan, P.K. and Majumder, H.K. (1998) A novel endonuclease from kinetoplastid hemo£agellated protozoan parasite Leishmania. J. Biochem. (Tokyo) 124, 1198^1205. Ho, H.-C., Shiau, P.-F., Liu, F.-C., Chung, J.-G. and Chen, L.-Y. (1998) Puri¢cation, characterization and complete amino acid sequence of nuclease C from Cunninghamella echinulata var. echinulata. Eur. J. Biochem. 256, 112^118. Nicieza, R.G., Huergo, J., Connolly, B.A. and Sanchez, J. (1999) Puri¢cation, characterization and role of nucleases and serine proteases in Streptomyces di¡erentiation. J. Biol. Chem. 274, 20366^ 20375. Rangarajan, E. and Shankar, V. (1999) Extracellular nuclease from Rhizopus stolonifer : puri¢cation and characteristics of single strand preferential deoxyribonuclease activity. Biochim. Biophys. Acta 1473, 293^304. Nomura, A., Suno, M. and Mizuno, Y. (1971) Studies on 3P-nucleotidase-nuclease from potato tubers I. Puri¢cation and some properties of the enzyme. J. Biochem. (Tokyo) 70, 993^1001. Imagawa, H., Toryu, H., Ozawa, T. and Takino, Y. (1982) Puri¢cation and characterization of nucleases from tea leaves. Agric. Biol. Chem. 46, 1261^1269. Matousek, J. and Tupy, J. (1984) Puri¢cation and properties of extracellular nuclease from tobacco pollen. Biol. Plant 26, 62^73. Brown, P.H. and Ho, T.-H.D. (1986) Barley aleurone layers secrete a nuclease in response to gibberillic acid. Plant Physiol. 82, 801^ 806. van der Westhuizen, A.J., Gliemeroth, A.K., Wenzel, W. and Hess, D. (1987) Isolation and partial characterization of an extracellular nuclease from pollen of Petunia hybrida. J. Plant Physiol. 131, 373^ 384. Kuligowska, E., Klarkowska, D. and Szarkowski, J.W. (1988) Puri¢cation and properties of endonuclease from wheat chloroplasts, speci¢c for single-stranded DNA. Phytochemistry 27, 1275^1279. Siwecka, M.A., Rytel, M. and Szarkowski, J.W. (1989) Puri¢cation and characterization of nuclease I associated with rye germ ribosomes. Acta Biochim. Pol. 36, 45^62. Siwecka, M.A. (1997) Puri¢cation and some properties of a novel dsRNA degrading nuclease bound to rye germ ribosomes. Acta Biochim. Pol. 44, 61^68. Vischi, M. and Marchetti, S. (1997) Strong extracellular nuclease activity displayed by barley (Hordeum vulgare L.) uninucleate microspores. Theor. Appl. Genet. 95, 185^190. Cordis, G.A., Goldblatt, P.-J. and Deutscher, M. (1975) Puri¢cation and characterization of a major endonuclease from rat liver nuclei. Biochemistry 14, 2596^2603. Ruiz-Carrillo, A. and Renaud, J. (1987) Endonuclease G: a (dG)nU(dC)n-speci¢c DNase from higher eukaryotes. EMBO J. 6, 401^407. Chou, M.-Y. and Liao, T.-H. (1990) Shrimp hepatopancreatic deoxyribonuclease ^ puri¢cation and characterization as well as comparison with bovine pancreatic deoxyribonuclease. Biochim. Biophys. Acta 1036, 95^100. Shuai, K., Das Gupta, C.K., Hawley, R.S., Chase, J.W., Stone, K.L. and Williams, K.R. (1992) Puri¢cation and characterization of an endo^exonuclease from adult £ies of Drosophila melanogaster. Nucleic Acids Res. 20, 1379^1385. Harosh, I., Mezzina, M., Harris, P.V. and Boyd, J.B. (1992) Puri¢cation and characterization of a mitochondrial endonuclease from Drosophila melanogaster embryos. Eur. J. Biochem. 210, 455^460. Tucker, P.W., Hazen Jr., E.E. and Cotton, F.A. (1978) Staphylococcal nuclease reviewed: A prototypic study in contemporary enzymol-

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

[56]

[57]

[58] [59] [60]

[61] [62] [63]

[64]

[65]

[66] [67] [68]

[69]

[70]

[71]

[72]

[73]

[74]

[75]

[76] [77]

[78] [79]

ogy. I. Isolation, physical and enzymatic properties. Mol. Cell. Biochem. 22, 67^77. Tucker, P.W., Hazen Jr., E.E. and Cotton, F.A. (1979) Staphylococcal nuclease reviewed : A prototypic study in contemporary enzymology. II. Solution studies of the nucleotide binding site and the e¡ects of nucleotide binding. Mol. Cell. Biochem. 23, 3^16. Tucker, P.W., Hazen Jr., E.E. and Cotton, F.A. (1979) Staphylococcal nuclease reviewed : A prototypic study in contemporary enzymology. III. Correlation of the three-dimensional structure with the mechanisms of enzymatic action. Mol. Cell. Biochem. 23, 67^86. Benedik, M.J. and Strych, U. (1998) Serratia marcescens and its extracellular nuclease. FEMS Microbiol. Lett. 165, 1^13. Martin, C.E. and Wagner, R.P. (1975) Two forms of a mitochondrial endonuclease in Neurospora crassa. Can. J. Biochem. 53, 823^825. Holstein, S.E.H., Kobert, B., Hillmer, S., Brown, P.H., Ho, T.-H.D. and Robinson, D.G. (1991) Subcellular localization of nuclease in barley aleurone. Physiol. Plant 83, 255^265. Coªte©, J. and Ruiz-Carillo, A. (1993) Primers for mitochondrial DNA replication generated by endonuclease G. Science 261, 765^769. Laskowski, M. and Seidel, M.D. (1945) Viscometric determination of thymonucleodepolymerase. Arch. Biochem. 7, 465^472. Schneider, W.C. and Hogeboom, G.H. (1952) Intracellular distribution of enzymes. X. Desoxyribonuclease and ribonuclease. J. Biol. Chem. 198, 155^163. Fujimoto, M., Fujiyama, K., Kuninaka, A. and Yoshino, H. (1974) Puri¢cation of a nuclease from Penicillium citrinum. Agric. Biol. Chem. 38, 777^783. Vogt, V.M. (1973) Puri¢cation and further properties of singlestrand-speci¢c nuclease from Aspergillus oryzae. Eur. J. Biochem. 33, 192^200. Roth, J.S. and Milstein, S.W. (1952) Ribonuclease I. A new method with P32 labeled yeast ribonucleic acid. J. Biol. Chem. 196, 489^500. Geiduschek, E.P. and Daniels, A. (1965) A simple assay for DNA endonucleases. Anal. Biochem. 11, 133^136. Kohen, R., Szyf, M. and Chevion, M. (1986) Quantitation of singleand double-strand DNA breaks in vitro and in vivo. Anal. Biochem. 154, 485^491. Pollack, Y., Kasir, J., Shemer, R., Metzger, S. and Szyf, M. (1984) Methylation pattern of mouse mitochondrial DNA. Nucleic Acids Res. 12, 4811^4824. Doetsch, P.W., McGray Jr., W.H. and Valenzuela, M.R.L. (1989) Partial puri¢cation and characterization of an endonuclease from spinach that cleaves ultraviolet light-damaged duplex DNA. Biochim. Biophys. Acta 1007, 309^317. Baumann, U., Frank, R. and Blo«cker, H. (1986) Conformational analysis of hairpin oligodeoxyribonucleotides by a single-strand-speci¢c nuclease. Eur. J. Biochem. 161, 409^413. Oleson, A.E. and Sasakuma, M. (1980) S1 nuclease of Aspergillus oryzae: A glycoprotein with an associated nucleotidase activity. Arch. Biochem. Biophys. 204, 361^370. Je¡ries, C.D., Holtman, D.F. and Guse, D.G. (1957) Rapid method for determining the activity of microorganisms on nucleic acids. J. Bacteriol. 73, 590^591. Horney, D.L. and Webster, D.A. (1971) Deoxyribonuclease : a sensitive assay using radial di¡usion in agarose containing methyl green^ DNA complex. Biochim. Biophys. Acta 247, 54^61. Boyd, J.B. and Mitchell, H.K. (1965) Identi¢cation of deoxyribonucleases in polyacrylamide gel following their separation by disk electrophoresis. Anal. Biochem. 13, 28^42. Rosenthal, A.L. and Lacks, S.A. (1977) Nuclease detection in SDS^ polyacrylamide gel electrophoresis. Anal. Biochem. 80, 76^90. Coªte©, J., Renaud, J. and Ruiz-Carillo, A. (1989) Recognition of (dG)n .(dC)n sequences by endonuclease G. Characterization of the calf thymus nuclease. J. Biol. Chem. 264, 3301^3310. Bernardi, G. (1973) Chromatography of proteins on hydroxyapatite. Methods Enzymol. 27, 471^479. Taniuchi, H. and Bohnert, J.L. (1975) The mechanism of stabilization

[80]

[81]

[82]

[83]

[84]

[85]

[86]

[87]

[88] [89]

[90]

[91]

[92]

[93]

[94]

[95]

[96]

[97]

609

of the structure of nuclease-T by binding of ligands. J. Biol. Chem. 250, 2388^2394. Janski, A.M. and Oleson, A.E. (1976) NADP-agarose : an a¤nity adsorbent for tobacco extracellular nuclease and other nucleases. Anal. Biochem. 71, 471^480. Gray, H.B. Jr., Winston, T.P., Hodnett, J.L., Legerski, R.J., Nees, D.W., Wei, C.-F. and Robberson, D.L. (1981) The extracellular nuclease from Alteromonas espejiana : An enzyme highly speci¢c for nonduplex structure in nominally duplex DNAs. In: Gene Ampli¢cation and Analysis, Vol. 2 (Chirikjian, J.G. and Papas, T.S., Eds.), pp. 169^203. Elsevier/North Holland, New York. Wei, C.-F., Alianell, G.A., Bencen, G.H. and Gray Jr., H.B. (1983) Isolation and comparison of two molecular species of the BAL 31 nuclease from Alteromonas espejiana with distinct kinetic properties. J. Biol. Chem. 258, 13506^13512. Taniuchi, H., An¢nsen, C.B. and Sodja, A. (1967) The amino acid sequence of an extracellular nuclease of Staphylococcus aureus. J. Biol. Chem. 242, 4752^4758. Friedho¡, P., Gimadutdinow, O., Ru«ter, T., Wende, W., Urbanke, C., Thole, H. and Pingoud, A. (1994) A procedure for renaturation and puri¢cation of the extracellular Serratia marcescens nuclease from genetically engineered Escherichia coli (Chem. Abstr. 120:185847). Protein Expr. Purif. 5, 37^43. Miller, M. and Krause, K. (1996) Identi¢cation of the Serratia endonuclease dimer : structural basis and implications for catalysis. Protein Sci. 5, 24^33. Franke, I., Meiss, G., Blecher, D., Gimadutdinow, O., Urbanke, C. and Pingoud, A. (1998) Genetic engineering, production and characterisation of monomeric variants of the dimeric Serratia marcescens endonuclease. FEBS Lett. 425, 517^522. Ball, T.K., Suh, Y. and Benedik, M.J. (1992) Disul¢de bonds are required for Serratia marcescens nuclease activity. Nucleic Acids Res. 20, 4971^4974. Rangarajan, E.S. (2001) Puri¢cation and characteristics of extracellular nuclease from Rhizopus stolonifer. Ph.D. dissertation, Pune. Franke, I., Meiss, G. and Pingoud, A. (1999) On the advantage of being a dimer, a case study using the dimeric Serratia nuclease and the monomeric nuclease from Anabaena sp. strain PCC 7120. J. Biol. Chem. 274, 825^832. Linn, S. and Lehman, I.R. (1965) An endonuclease from Neurospora crassa speci¢c for polynucleotides lacking an ordered structure I. Puri¢cation and properties of the enzyme. J. Biol. Chem. 240, 1287^1293. Linn, S. and Lehman, I.R. (1965) An endonuclease from Neurospora crassa speci¢c for polynucleotides lacking an ordered structure II. Studies on enzyme speci¢city. J. Biol. Chem. 240, 1294^1304. Fraser, M.J., Tjeerde, R. and Matsumoto, K. (1976) A second form of the single-strand speci¢c endonuclease of Neurospora crassa which is associated with a double-strand exonuclease. Can. J. Biochem. 54, 971^980. Fraser, M.J., Chow, T.Y.-K., Cohen, H. and Koa, H. (1986) An immunochemical study of Neurospora crassa nucleases. Biochem. Cell Biol. 64, 106^116. Kwong, S. and Fraser, M.J. (1978) Neurospora crassa endoexonuclease and its inactive (precursor?) form. Can. J. Biochem. 56, 370^ 377. Chesbro, W.R., Stuart, D. and Burke II, J.J. (1966) Multiple molecular weight forms of staphylococcal nuclease. Biochem. Biophys. Res. Commun. 23, 783^792. Filimonova, M.N., Dementyev, A.A., Leshchinskaya, I.B., Bakulina, G.Yu. and Shlyapnikov, S.V. (1991) Isolation and characterization of extracellular nuclease of Serratia marcescens. Biokhimiya 50, 508^ 520. Pedersen, J., Andersen, J., Roepstor¡, P., Filimonova, M. and Biedermann, K. (1993) Characterization of natural and recombinant nuclease isoforms by electrospray mass spectrometry. Biotechnol. Appl. Biochem. 18, 389^399.

FEMSRE 729 5-12-01

610

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

[98] Pedersen, J., Filimonova, M., Roepstro¡, P. and Biedermann, K. (1993) Characterization of Serratia marcescens nuclease isoforms by plasma desorption mass spectrometry. Biochim. Biophys. Acta 1202, 13^21. [99] Taniuchi, H. and An¢nsen, C.B. (1968) Steps in the formation of active derivatives of staphylococcal nuclease during trypsin digestion. J. Biol. Chem. 243, 4778^4786. [100] Lin, J.-L., Wang, W.-Y. and Liao, T.-H. (1994) Thermal inactivation of shrimp deoxyribonuclease with and without sodium dodecyl sulfate. Biochim. Biophys. Acta 1209, 209^214. [101] Heins, J.N., Suriano, J.R., Taniuchi, H. and An¢nsen, C.B. (1967) Characterization of a nuclease produced by Staphylococcus aureus. J. Biol. Chem. 242, 1016^1020. [102] Brown, P.H. and Ho, T.-H.D. (1987) Biochemical properties and hormonal regulation of barley nuclease. Eur. J. Biochem. 168, 357^ 364. [103] Shimada, H., Inokuchi, N., Koyama, T. and Irie, M. (1991) Puri¢cation and characterization of a nuclease from Lentinus edodes. Chem. Pharm. Bull. 39, 2633^2637. [104] Kobayashi, H., Inokuchi, N., Koyama, T., Tomita, M. and Irie, M. (1995) Puri¢cation and characterization of the 2nd 5P-nucleotideforming nuclease from Lentinus edodes. Biosci. Biotechnol. Biochem. 59, 1169^1171. [105] Cuatrecasas, P., Fuchs, S. and An¢nsen, C.B. (1967) Catalytic properties and speci¢city of the extracellular nuclease of Staphylococcus aureus. J. Biol. Chem. 242, 1541^1547. [106] Rangarajan, E.S. and Shankar, V. (2001) Nuclease Rsn from Rhizopus stolonifer : characteristics of associated ^ adenine speci¢c ^ ribonuclease activity. J. Biochem. Mol. Biol. Biophys. 5, 99^108. [107] Fraser, M.J. (1980) Puri¢cation and properties of Neurospora crassa endo^exonuclease, an enzyme which can be converted to a singlestrand speci¢c endonuclease. Methods Enzymol. 65, 255^263. [108] von Hippel, P.H. and Felsenfeld, G. (1964) Micrococcal nuclease as a probe of DNA conformation. Biochemistry 3, 27^35. [109] Wingert, L. and von Hippel, P.H. (1968) The conformation dependent hydrolysis of DNA by micrococcal nuclease. Biochim. Biophys. Acta 157, 114^126. [110] Nestle, M. and Roberts, W.K. (1969) An extracellular nuclease from Serratia marcescens I. Puri¢cation and some properties of the enzyme. J. Biol. Chem. 244, 5213^5218. [111] Filimonova, M.N., Balaban, N.P., Sharipova, F.R. and Leshchinskaya, I.B. (1980) Production of Serratia marcescens nuclease in a homogeneous state and study of the physicochemical properties of the enzyme. Biokhimiya 45, 2097^2103. [112] Shishido, K. (1985) E¡ect of spermine on cleavage of plasmid DNA by nuclease S1 and BAL 31. Biochim. Biophys. Acta 826, 147^ 150. [113] Frank, J.J., Hawk, J.A. and Levy, C.C. (1975) Polyamine activation of staphylococcal nuclease. Biochim. Biophys. Acta 390, 117^124. [114] Levy, C.C., Hieter, P.A. and LeGendre, S.M. (1974) Evidence for the direct binding of polyamines to a ribonuclease that hydrolyzes ribonucleic acid at uridylic acid residues. J. Biol. Chem. 249, 6762^ 6769. [115] Filimonova, M.N., Baratova, L.A., Vospel'nikova, M.D., Zheltova, A.O. and Leshchinskaya, I.B. (1981) Characterization of endonuclease from Serratia marcescens. Biokhimiya 46, 1660^1666. [116] Suno, M., Nomura, A. and Mizuno, Y. (1973) Studies on 3P-nucleotidase-nuclease from potato tubers II. Further studies on substrate speci¢city and mode of action. J. Biochem. (Tokyo) 73, 1291^1297. [117] Hatahet, Z. and Fraser, M.J. (1989) Speci¢c inhibitors of Neurospora endo^exonuclease. Biochem. Cell Biol. 67, 632^641. [118] Meiss, G., Franke, I., Gimadutdinow, O., Urbanke, C. and Pingoud, A. (1998) Biochemical characterization of Anabaena sp. strain PCC 7120 non-speci¢c nuclease NucA and its inhibitor NuiA. Eur. J. Biochem. 251, 924^934. [119] Meiss, G., Gimadutdinow, O., Haberland, B. and Pingoud, A. (2000) Mechanism of DNA cleavage by the DNA/RNA-non-speci¢c

[120]

[121]

[122]

[123]

[124]

[125]

[126]

[127]

[128] [129]

[130]

[131]

[132]

[133]

[134]

[135]

[136]

[137]

[138]

[139] [140]

[141]

Anabaena sp. PCC 7120 endonuclease NucA and its inhibtion by NuiA. J. Mol. Biol. 297, 521^534. Chen, Y.-C., Suh, Y., Riise, E., Kartman, B., Jin, S. and Benedik, M.J. (1996) Inhibition of Serratia marcescens nuclease secretion by a truncated nuclease peptide. Gene 172, 9^16. Oleson, A.E. and Hoganson, E.D. (1981) S1 nuclease of Aspergillus oryzae: characterization of the associated phosphomonoesterase activity. Arch. Biochem. Biophys. 211, 478^484. Rusche, J.R., Rowe, T.C. and Holloman, W.K. (1980) Puri¢cation and characterization of nuclease L from Ustilago maydis. J. Biol. Chem. 255, 9117^9123. Desai, N.A. and Shankar, V. (2000) Puri¢cation and characterization of the single-strand-speci¢c and guanylic-acid-preferential deoxyribonuclease activity of the extracellular nuclease from Basidiobolus haptosporus. Eur. J. Biochem. 267, 5123^5135. Sulkowski, E. and Laskowski Sr., M. (1968) The occurrence of autoacceleration during hydrolysis of nucleic acids by micrococcal nuclease. J. Biol. Chem. 243, 4917^4921. Stevens, A. and Hilmoe, R.J. (1960) Studies on a nuclease from Azotobacter agilis II. Hydrolysis of ribonucleic and deoxyribonucleic acid. J. Biol. Chem. 235, 3023^3027. Nestle, M. and Roberts, W.K. (1969) An extracellular nuclease from Serratia marcescens II. Speci¢city of the enzyme. J. Biol. Chem. 244, 5219^5225. Khorana, H.G. (1961) Phosphodiesterases. In: The Enzymes, Vol. 5 (Boyer, P.D., Lardy, H. and Myrback, K., Eds.), pp. 79^94. Academic Press, New York. Razzell, W.E. (1963) Phosphodiesterases. Methods Enzymol. 6, 236^ 258. Razzell, W.E. and Khorana, H.G. (1959) Studies on polynucleotides III. Enzymic degradation. Substrate speci¢city and properties of snake venom phosphodiesterase. J. Biol. Chem. 234, 2105^2113. Bernardi, A. and Bernardi, G. (1968) Studies on acid hydrolases. V. Isolation and characterization of spleen nucleoside polyphosphatase. Biochim. Biophys. Acta 155, 371^377. Liao, T.-H. (1975) Deoxythymidine 3P,5P-di-p-nitrophenyl phosphate as a synthetic substrate for bovine pancreatic deoxyribonuclease. J. Biol. Chem. 250, 3721^3724. Cuatrecasas, P., Wilchek, M. and An¢nsen, C.B. (1969) The action of staphylococcal nuclease on synthetic substrates. Biochemistry 8, 2277^2284. Friedho¡, P., Meiss, G., Kolmes, B., Pieper, U., Gimadutdinow, O., Urbanke, C. and Pingoud, A. (1996) Kinetic analysis of the cleavage of natural and synthetic substrates by the Serratia nuclease. Eur. J. Biochem. 241, 572^580. Kolmes, B., Franke, I., Friedho¡, P. and Pingoud, A. (1996) Analysis of the reaction mechanism of the non-speci¢c endonuclease of Serratia marcescens using an arti¢cial minimal substrate. FEBS Lett. 397, 343^346. Pritchard, A.E., Kowalski, D. and Laskowski Sr., M. (1977) An endonuclease activity of venom phosphodiesterase speci¢c for single-stranded and superhelical DNA. J. Biol. Chem. 252, 8652^8659. Williams, E.J., Sung, S.-C. and Laskowski Sr., M. (1961) Action of venom phosphodiesterase on deoxyribonucleic acid. J. Biol. Chem. 236, 1130^1134. Razzell, W.E. and Khorana, H.G. (1961) Studies on polynucleotides X. Enzymic degradation. Some properties and mode of action of spleen phosphodiesterase. J. Biol. Chem. 236, 1144^1149. Ito, K., Matsuura, Y. and Minamiura, N. (1994) Puri¢cation and characterization of fungal nuclease composed of heterogeneous subunits. Arch. Biochem. Biophys. 309, 160^167. Thomas Jr., C.A. (1956) The enzymatic degradation of desoxyribose nucleic acid. J. Am. Chem. Soc. 78, 1861^1868. Schumaker, V.N., Richards, E.G. and Schachman, H.K. (1956) A study of the kinetics of the enzymatic digestion of deoxyribonucleic acid. J. Am. Chem. Soc. 78, 4230^4236. Bernardi, G. and Cordonnier, C. (1965) Mechanism of degradation

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613

[142]

[143]

[144]

[145] [146]

[147]

[148]

[149]

[150]

[151]

[152]

[153]

[154]

[155]

[156]

[157]

[158]

[159]

[160] [161]

[162]

of DNA by endonuclease I from Escherichia coli. J. Mol. Biol. 11, 141^143. Weir, A.F. (1993) Deoxyribonuclease I (EC 3.1.21.1) and II (EC 3.1.22.1). In: Methods in Molecular Biology, Vol. 16 (Burell, M.M., Ed.), pp. 7^16. Humana Press, Totowa, NJ. Melgar, E. and Goldthwait, D.A. (1968) Deoxyribonucleic acid nucleases: The e¡ects of metals on the mechanism of action of deoxyribonuclease I. J. Biol. Chem. 243, 4409^4416. Meiss, G., Friedho¡, P., Hahn, M., Gimadutdinow, O. and Pingoud, A. (1995) Sequence preferences in cleavage of dsDNA and ssDNA by the extracellular Serratia marcescens endonuclease. Biochemistry 34, 11979^11988. Zhou, X. and Gray Jr., H.B. (1990) Mechanism of exonuclease action of BAL 31 nuclease. Biochim. Biophys. Acta 1049, 83^91. Gray, Jr. H.B. and Lu, T. (1993) The BAL 31 nucleases (EC 3.1.11). In: Methods in Molecular Biology, Vol. 16 (Burrel, M.M., Ed.), pp. 231^251. Humana Press, Totowa, NJ. Bencen, G.H., Wei, C.-F., Robberson, D.L. and Gray Jr., H.B. (1984) Terminally directed hydrolysis of duplex ribonucleic acid catalysed by a species of the BAL 31 nuclease from Alteromonas espejiana. J. Biol. Chem. 259, 13584^13589. Lu, T. and Gray Jr., H.B. (1995) Kinetics and mechanism of BAL 31 nuclease action on small substrates and single-stranded DNA. Biochim. Biophys. Acta 1251, 125^138. Kanamori, N., Cozzarelli, N.R. and Okazaki, R. (1974) Extracellular nucleases of Bacillus subtilis II. The nucleases as 5P-end-group reagents. Biochim. Biophys. Acta 335, 173^184. Bauer, W. and Vinograd, J. (1974) Circular DNAs. In: Basic Principles of Nucleic Acid Chemistry, Vol. 2 (Cantoni, G.L. and Davis, D.R., Eds.), pp. 265^303. Academic Press, New York. Przykorska, A.K., Hauser, C.R. and Gray Jr., H.B. (1988) Circular intermediates with missing nucleotides in the conversion of supercoiled or nicked circular to linear duplex DNA catalyzed by two species of BAL 31 nuclease. Biochim. Biophys. Acta 949, 16^26. Sakaguchi, R., Joho, K. and Shishido, K. (1985) E¡ect of netropsin on plasmid DNA cleavage by BAL 31 nuclease. FEBS Lett. 191, 59^ 62. Wei, C.-F., Legerski, R.J., Alianell, G.A., Robberson, D.L. and Gray Jr., H.B. (1984) A single apurinic site can elicit BAL 31 nuclease-catalyzed cleavage of duplex DNA. Biochim. Biophys. Acta 782, 408^414. Dirksen, M.L. and Dekker, C.A. (1960) Micrococcal nuclease: Consideration of its mode of action. Biochem. Biophys. Res. Commun. 2, 147^152. Felsenfeld, G. and Sandeen, G. (1962) The dispersion of the hyperchromic e¡ect in thermally induced transitions of nucleic acids. J. Mol. Biol. 5, 587^610. Galcheva-Gargova, Z., Davidov, V. and Dessev, G. (1985) Formation of single-stranded regions in the course of digestion of DNA with DNase II and micrococcal nuclease. Arch. Biochem. Biophys. 240, 464^469. Meiss, G., Gast, F.-U. and Pingoud, A.M. (1999) The DNA/RNA non-speci¢c Serratia nuclease prefers double-stranded A-form nucleic acids as substrates. J. Mol. Biol. 288, 377^390. Arnott, S. and Selsing, E. (1974) Letter: The structure of polydeoxyguanylic acid with polydeoxycytidylic acid. J. Mol. Biol. 88, 551^ 552. McCall, M., Brown, T. and Kennard, O. (1985) The crystal structure of d(G-G-G-G-C-C-C-C). A model for poly(dG)Upoly(dC). J. Mol. Biol. 183, 385^396. Zimmermann, S.B. (1982) The three-dimensional structure of DNA. Annu. Rev. Biochem. 51, 395^427. Kilpatrick, M.W., Wei, C.F., Gray Jr., H.B. and Wells, R.D. (1983) BAL 31 nuclease as a probe in concentrated salt for the B^Z DNA junction. Nucleic Acids Res. 11, 3811^3822. Filimonova, M.N., Garusov, A.V., Smetanina, T.A., Andreeva, M.A., Bogomolnaya, L. and Leshchinskaya, I.B. (1996) Isoforms

[163]

[164]

[165]

[166]

[167]

[168]

[169] [170]

[171]

[172]

[173]

[174]

[175]

[176]

[177]

[178]

[179]

[180]

611

of Serratia marcescens nuclease. Comparative analysis of substrate speci¢city. Biochemistry (Moscow) 61, 1274^1278. Pedersen, J., Filimonova, M.N., Roepstor¡, P. and Biedermann, K. (1995) Isoforms of Serratia marcescens nuclease produced by natural and recombinant strains. Comparative characterization by plasma desorption mass spectrometry. Biokhimiya 60, 450^461. Mikulski, A.J., Sulkowski, E., Stasiuk, L. and Laskowski Sr., M. (1969) Susceptibility of dinucleotides bearing either 3P or 5P-monophosphate to micrococcal nuclease. J. Biol. Chem. 244, 6559^6565. Vincent, R.D., Hofmann, T.J. and Zassenhaus, J.P. (1988) Sequence and expression of NUC1, the gene encoding the mitochondrial nuclease in Saccharomyces cerevisiae. Nucleic Acids Res. 16, 3297^ 3312. Friedho¡, P., Gimadutdinow, O. and Pingoud, A. (1994) Identi¢cation of catalytically relevant amino acids of the extracellular Serratia marcescens endonuclease by alignment-guided mutagenesis. Nucleic Acids Res. 22, 3280^3287. Friedho¡, P., Kolmes, B., Gimadutdinow, O., Wende, W., Krause, K.L. and Pingoud, A. (1996) Analysis of the mechanism of the Serratia nuclease using site-directed mutagenesis. Nucleic Acids Res. 24, 2632^2639. Wang, W.-Y., Liaw, S.-H. and Liao, T.-H. (2000) Cloning and characterization of a novel nuclease from shrimp hepatopancreas, and prediction of its active site. Biochem. J. 346, 799^804. Ho, H.-C. and Liao, T.-H. (1999) Protein structure and gene cloning of Syncephalastrum racemosum nuclease. Biochem. J. 339, 261^267. Ball, T.K., Saurugger, P.N. and Benedik, M.J. (1987) The extracellular nuclease gene of Serratia marcescens and its secretion from Escherichia coli. Gene 57, 183^192. Biedermann, K., Jepsen, P.K., Riise, E. and Svendsen, I. (1989) Puri¢cation and characterization of a Serratia marcescens nuclease produced by Escherichia coli (Chem. Abstr. 112:174481). Carlsberg Res. Commun. 54, 17^27. Bannikova, G.E., Blagova, E.V., Dementiev, A.A., Morgunova, E.Y., Mikchailov, A.M., Shlyapnikov, S.V., Varlamov, V.P. and Vainshtein, B.K. (1991) Two isoforms of Serratia marcescens nuclease. Crystallization and preliminary X-ray investigation of the enzyme. Biochem. Mol. Biol. Int. 24, 813^822. Pedersen, J., Filimonova, M.N., Roepstor¡, P. and Biedermann, K. (1995) Serratia marcescens nucleases. Part II. Analysis of primary structures by peptide mapping in combination with plasma desorption mass spectrometry. Bioorg. Khim. 21, 336^344. Arnone, A., Bier, C.J., Cotton, F.A., Day, V.W., Hazen Jr., E.E., Richardson, D.C., Richardson, J.S. and Yonath, A. (1971) A high resolution structure of an inhibitor complex of the extracellular nuclease of Staphylococcus aureus. J. Biol. Chem. 246, 2302^2316. Cotton, F.A., Hazen, E.A. and Legg, M.J. (1979) Staphylococcal nuclease: proposed mechanism of action based on structure of enî zyme^thymidine 3P,5P-bisphosphate^calcium ion complex at 1.5-A resolution. Proc. Natl. Acad. Sci. USA 76, 2551^2555. Loll, P.J. and Lattman, E.E. (1989) The crystal structure of the ternary complex of staphylococcal nuclease, Ca2‡ , and the inhibitor î . Proteins Struct. Funct. Genet. 5, 183^201. pdTp, re¢ned at 1.65 A Miller, M.D., Tanner, J., Alpaugh, M., Benedik, M.J. and Krause, î structure of Serratia endonuclease suggests a K.L. (1994) 2.1 A mechanism for binding to double-stranded DNA. Nat. Struct. Biol. 1, 461^468. Lunin, V.Y., Levdikov, V.M., Shlyapnikov, S.V., Blagova, E.V., Lunin, V.V., Wilson, K.S. and Mikhailov, A.M. (1997) Three-diî resomensional structure of Serratia marcescens nuclease at 1.7 A lution and mechanism of its action. FEBS Lett. 412, 217^222. Shlyapnikov, S.V., Lunin, V.V., Perbandt, M., Polyakov, K.M., Lunin, V.Y., Levdikov, V.M., Betzel, C. and Mikhailov, A.M. (2000) Atomic structure of the Serratia marcescens endonuclease î resolution and the enzyme reaction mechanism. Acta Crysat 1.1 A tallogr. D 56, 567^572. Cuatrecasas, P., Fuchs, S. and An¢nsen, C.B. (1967) The binding of

FEMSRE 729 5-12-01

612

[181]

[182]

[183]

[184]

[185]

[186]

[187]

[188]

[189]

[190]

[191]

[192]

[193]

[194]

[195]

[196]

[197]

[198]

[199]

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613 nucleotides and calcium to the extracellular nuclease of Staphylococcus aureus. Studies by gel ¢ltration. J. Biol. Chem. 242, 3063^3067. Cuatrecasas, P., Fuchs, S. and An¢nsen, C.B. (1969) Cross-linking of aminotyrosyl residues in the active site of staphylococcal nuclease. J. Biol. Chem. 244, 406^412. Uhlmann, E. and Smith, J.A. (1987) Site-directed mutagenesis of staphylococcal nuclease: Role of tyrosine residues in substrate binding and catalysis. Nucleosides Nucleotides 6, 331^334. Miller, M.D., Cai, J. and Krause, K.L. (1999) The active site of Serratia endonuclease contains a conserved magnesium^water cluster. J. Mol. Biol. 288, 975^987. Rangarajan, E.S. and Shankar, V. (2000) Active site characterization of nuclease Rsn from Rhizopus stolonifer : involvement of histidine in catalysis, tryptophan in substrate binding and carboxylate in metal binding. J. Biochem. Mol. Biol. Biophys. (in press). Cotton, F.A. and Hazen, Jr. E.E. (1971) Staphylococcal nuclease. X-ray structure. In: The Enzymes, Vol. 3 (Boyer, P.D., Ed.), pp. 153^170. Academic Press, New York. Serpersu, E.H., Shortle, D. and Mildvan, A.S. (1987) Kinetic and magnetic resonance studies of active-site mutants of staphylococcal nuclease: factors contributing to catalysis. Biochemistry 26, 1289^ 1300. Libson, A.M., Gittis, A.G. and Lattman, A.E. (1994) Crystal structures of the binary Ca2‡ and pdTp complexes and the ternary complex of the Asp21CGlu mutant of staphylococcal nuclease. Implications for catalysis and ligand binding. Biochemistry 33, 8007^ 8016. Cuatrecasas, P., Edelhoch, H. and An¢nsen, C.B. (1967) Fluorescence studies of the interaction of nucleotides with the active site of the nuclease of Staphylococcus aureus. Proc. Natl. Acad. Sci. USA 58, 2043^2050. Cuatrecasas, P., Wilchek, M. and An¢nsen, C.B. (1968) Staphylococcal nuclease: Size and speci¢city of the active site. Science 162, 1491^1493. Weber, D.J., Mullen, G.P. and Mildvan, A.S. (1991) Conformation of an enzyme-bound substrate of staphylococcal nuclease as determined by NMR. Biochemistry 30, 7425^7437. Stanczyk, S.M. and Bolton, P.M. (1992) Comparison of conformational features of staphylococcal nuclease in ternary complexes with pdTp, pdGp, and nitrophenyl-pdTp. Biochemistry 31, 6396^6401. Grissom, C.B. and Markley, J.L. (1989) Staphylococcal nuclease active-site amino acids: pH dependence of tyrosines and arginines by 13 C NMR and correlation with kinetic studies. Biochemistry 28, 2116^2124. L'vova, T.H., Avrosimova-amel-ianchik, N.M. and Tatarskaia, R.I. (1976) A mechanism of mononucleotide formation under endonuclease hydrolysis. Biokimiya 41, 2077^2081. Srivastava, T.K., Friedho¡, P., Pingoud, A. and Katti, S.B. (1999) Application of oligonucleoside methylphosphonates in the studies on phosphodiester hydrolysis by Serratia endonuclease. Nucleosides Nucleotides 18, 1945^1960. An¢nsen, C.B., Cuatrecasas, P. and Taniuchi, H. (1971) Staphylococcal nuclease. Chemical properties and catalysis. In: The Enzymes, 3rd edn., Vol. 4 (Boyer, P.D., Ed.), pp. 177^204. Academic Press, New York. Chaiken, I.M. and Sanchez, G.R. (1972) Active site properties of (aspartic acid 43)-semisynthetic nuclease-TP. J. Biol. Chem. 247, 6743^6747. Serpersu, E.H., Shortle, D. and Mildvan, A.S. (1986) Kinetic and magnetic resonance studies of e¡ects of genetic substitution of a Ca2+-liganding amino acid in staphylococcal nuclease. Biochemistry 25, 68^77. Serpersu, E.H., McCracken, J., Peisach, J. and Mildvan, A.S. (1988) Electron spin echo modulation and nuclear relaxation studies of staphylococcal nuclease and its metal-coordinating mutants. Biochemistry 27, 8034^8044. Serpersu, E.H., Hibler, E.W., Gerlt, J.A. and Mildvan, A.S. (1989)

[200]

[201]

[202]

[203]

[204]

[205]

[206]

[207]

[208]

[209]

[210]

[211]

[212]

[213]

[214]

[215] [216]

[217]

Kinetic and magnetic resonance studies of the glutamate-43 to serine mutant of staphylococcal nuclease. Biochemistry 28, 1539^1548. Hibler, D.W., Stolowich, J.N., Reynolds, M.A., Gerlt, J.A., Wilde, J.A. and Bolton, P.H. (1987) Site-directed mutants of staphylococcal nuclease. Detection and localization by proton NMR spectroscopy of conformational changes accompanying substitutions for glutamic acid-43. Biochemistry 26, 6278^6286. Weber, D.J., Serpersu, E.H., Shortle, D. and Mildvan, A.S. (1990) Diverse interactions between the individual mutations in a double mutant at the active site of staphylococcal nuclease. Biochemistry 29, 8632^8642. Ste¡ens, J.J., Siewers, I.J. and Benkovic, S.J. (1975) Catalysis of phosphoryl group transfer. The role of divalent metal ions in the hydrolysis of lactic acid O-phenyl phosphate and salicylic acid Oaryl phosphate. Biochemistry 14, 2431^2440. Judice, J.K., Gamble, T.R., Murphy, E.C., de Vos, A.M. and Schultz, P.G. (1993) Probing the mechanism of staphylococcal nuclease with unnatural amino acids : Kinetic and structural studies. Science 261, 1578^1581. Loll, P.J., Quirk, S., Lattman, E.E. and Garavito, R.M. (1995) Xray crystal structures of staphylococcal nuclease complexed with the competitive inhibitor cobalt(II) and nucleotide. Biochemistry 34, 4316^4324. Loll, P.J. and Lattman, E.E. (1990) Active site mutant Glu-43CAsp in staphylococcal nuclease displays nonlocal structural changes. Biochemistry 29, 6866^6873. Mehdi, S. and Gerlt, J.A. (1982) Oxygen chiral phosphodiesters. 7. Stereochemical course of a reaction catalyzed by staphylococcal nuclease. J. Am. Chem. Soc. 104, 3223^3225. Potter, B.V.L., Connolly, B.A. and Eckstein, F. (1983) Synthesis and con¢gurational analysis of a dinucleoside phosphate isotopically chiral at phosphorus. Stereochemical course of Penicillium citrinum nuclease P1 reaction. Biochemistry 22, 1369^1377. Potter, B.V.L., Romaniuk, P.J. and Eckstein, F. (1983) Stereochemical course of DNA hydrolysis by nuclease S1. J. Biol. Chem. 258, 1758^1760. Hale, S.P., Poole, L.B. and Gerlt, J.A. (1993) Mechanism of the reaction catalyzed by staphylococcal nuclease: identi¢cation of the rate-determining step. Biochemistry 32, 7479^7487. Friedho¡, P., Franke, I., Krause, K.L. and Pingoud, A. (1999) Cleavage experiments with deoxythymidine 3P,5P-bis-(p-nitrophenyl phosphate) suggest that the homing endonuclease I-PpoI follows the same mechanism of phosphodiester bond hydrolysis as the non-speci¢c Serratia nuclease. FEBS Lett. 443, 209^214. Friedho¡, P., Franke, I., Meiss, G., Wende, W., Krause, K.L. and Pingoud, A. (1999) A similar active site for non-speci¢c endonucleases. Nat. Struct. Biol. 6, 132^133. Ku«hlmann, U.C., Moore, C.R., James, R., Kleanthous, C. and Hemmings, A.M. (1999) Structural parsimony in endonuclease active sites: should the number of homing endonuclease families be rede¢ned? FEBS Lett. 463, 1^2. Raaijmakers, H., Toro, I., Birkenbihl, R., Kemper, B. and Suck, D. (2001) Conformational £exibility in T4 endonuclease VII revealed by crystallography : implications for substrate binding and cleavage. J. Mol. Biol. 308, 311^323. Romier, C., Domingues, R., Lahm, A., Dahl, O. and Suck, D. (1998) Recognition of single-stranded DNA by nuclease P1: High resolution crystal structures of complexes with substrate analogs. Proteins Struct. Funct. Genet. 32, 414^424. Wilcox, D.E. (1996) Binuclear metallohydrolases. Chem. Rev. 96, 2435^2458. Fraser, M.J. (1979) Alkaline deoxyribonucleases released from Neurospora crassa mycelia : two activities not released by mutants with multiple sensitivities to mutagens. Nucleic Acids Res. 6, 231^246. Hess, D. (1975) Uptake of DNA and bacteriophage into pollen and genetic manipulation. In: Genetic Manipulations with Plant Material (Ledoux, L., Ed.), pp. 519^537. Plenum Press, New York.

FEMSRE 729 5-12-01

E.S. Rangarajan, V. Shankar / FEMS Microbiology Reviews 25 (2001) 583^613 [218] Hess, D. (1987) Pollen based techniques in genetic manipulation. In: Pollen Development and Cytology (Giles, K.L. and Prakash, J., Eds.), pp. 367^395. Academic Press, New York. [219] Compton, M.M. and Cidlowski, J.A. (1987) Identi¢cation of a glucocorticoid-induced nuclease in thymocytes. A potential `lysis gene' product. J. Biol. Chem. 262, 8288^8292. [220] Gaido, M.L. and Cidlowski, J.A. (1991) Identi¢cation, puri¢cation, and characterization of a calcium-dependent endonuclease (NUC18) from apoptotic rat thymocytes. NUC18 is not histone H2B. J. Biol. Chem. 266, 18580^18585. [221] Montague, J.W., Gaido, M.L., Frye, C.W. and Cidlowski, J.A. (1994) A calcium-dependent nuclease from apoptotic rat thymocytes is homologous with cyclophilin. Recombinant cyclophilins A, B, and C have nuclease activity. J. Biol. Chem. 269, 18877^18880. [222] Montague, J.W., Hughes Jr., F.M. and Cidlowski, J.A. (1997) Native recombinant cyclophilins A, B, and C degrade DNA independently of peptidylprolyl cis-trans-isomerase activity. Potential roles of cyclophilins in apoptosis. J. Biol. Chem. 272, 6677^6684. [223] Sachs, D.H., Berzofsky, J.A., Pisetsky, D.S. and Schwartz, R.H. (1978) Genetic control of the immune response to staphylococcal nuclease (Chem. Abstr. 89:105550). Springer Semin. Immunopathol. 1, 51^83. [224] Gudding, R. (1980) Measurement of Staphylococcus aureus nuclease and antinucleases. Applicability for the assessment of mastitic milk (Chem. Abstr. 93:163241). Acta Vet. Scand. 21, 229^241. [225] Gudding, R. (1980) Nuclease of Staphylococcus epidermidis isolated from mastitic milk. Production and some properties (Chem. Abstr. 93:110305). Acta Vet. Scand. 21, 267^277. [226] Sundaram, S.P., Kumari, S.L.S. and Murthy, K.V. (1982) Thermostable nuclease activity of Staphylococci from clinical sources (Chem. Abstr. 96:177612). Indian J. Med. Res. 75, 19^22.

613

[227] Stersky, A.K., Szabo, R., Todd, E.C.D., Thacker, C., Dickie, N. and Akhtar, M. (1986) Staphylococcus aureus growth and thermostable nuclease and enterotoxin production in canned salmon and sardines (Chem. Abstr. 105:77691). J. Food Prot. 49, 428^435. [228] Nunez, M., Bautista, L., Medina, M. and Gaya, P. (1988) Staphylococcus aureus, thermostable nuclease and staphylococcal enterotoxins in raw ewes' milk Manchego cheese (Chem. Abstr. 110:6595). J. Appl. Bacteriol. 65, 29^34. [229] Schiebel, E., Schwarz, H. and Braun, V. (1989) Subcellular location and unique secretion of the hemolysin of Serratia marcescens. J. Biol. Chem. 264, 16311^16320. [230] Hejazi, A. and Falkiner, F.R. (1997) Serratia marcescens (Chem. Abstr. 128:20317). J. Med. Microbiol. 46, 903^912. [231] Matousek, J., Trnena, L., Oberhauser, R., Lichtenstein, C.P. and Nellen, W. (1994) dsRNA degrading nucleases are di¡erentially expressed in tobacco anthers. Biol. Chem. Hoppe-Seyler 375, 261^ 269. [232] Shapiro, J., Machattie, L., Eron, L., Ihler, G., Ippen, K. and Beckwith, J. (1969) Isolation of pure lac operon DNA. Nature (London) 224, 768^774. [233] Marks, A. and Spencer, J.H. (1970) Isolation of Escherichia coli transfer RNA hybrids. J. Mol. Biol. 51, 115^130. [234] Joseph, D.R. and Sta¡ord, D.W. (1976) Puri¢cation of sea urchin ribosomal RNA genes with a single-strand speci¢c nuclease. Biochim. Biophys. Acta 418, 167^174. [235] Bartok, K., Fraser, M.J. and Fareed, G.C. (1974) Detection of sequence heterology by use of Neurospora crassa nucleases. Biochem. Biophys. Res. Commun. 60, 507^514. [236] Krupp, G. and Gross, H.J. (1979) Rapid RNA sequencing: nucleases from Staphylococcus aureus and Neurospora crassa discriminate between uridine and cytidine. Nucleic Acids Res. 6, 3481^3490.

FEMSRE 729 5-12-01