Volume 1, 2016
 
 
Archive
 
+ Volume 1 2016
Issue 1
 
 
Insight Medicine>>  Volume 1 Issue 1, 2016
Download  |  PDF  

An Overview on Ribonucleases and their Therapeutic Effects
Gupta Shruti , Singh Sukhdev and Kanwar Shamsher Singh
 
ABSTRACT:
The maturation and degradation of ribonucleic acid (RNA) into smaller nucleotides are carried out by a wide variety of cellular ribonucleases (RNases) which are also responsible for regulating the functional expression of several fundamental genes in living systems. They exist in eukaryotes to prokaryotes as well as in viruses. Structure, functions and catalytic mechanism of RNases have been well understood especially in Escherichia coli model. In addition to regulate the RNA metabolisms RNases demonstrate wide range of therapeutic effects also. They exhibit antitumor, antifungal, antiviral and immunosuppressive functions. Bovine pancreatic RNase (RNase A), bovine seminal RNase (BS-RNase), onconase and angiogenin are significant antitumor RNases. Onconase (Rampinase) is a stable amphibian homolog which displays significant cytotoxicity towards tumor cells and is the only ribonuclease which has reached up to the clinical levels. Onconase has proved potential therapeutic applications in the treatment of prostate, cervical, colon, breast and ovarian cancers as well as in lymphocytic leukaemia. The RNases demonstrate antiviral properties also. The Eosinophil Derived Neurotoxin (EDN) and the Eosinophil Cationic Protein (ECP) also known as RNases 2 and 3 are such RNases. The EDN shows broad range of antiviral activity towards several RNA viruses including HIV-1. Recombinant EDN constructs for example EDNsFv and rhEDN express cytotoxic effects against transfer receptor(s) expressing leukemia cells and respiratory syncytial virus. Some other RNases i.e., RNase 5, RNase 7 and RNase 8 have also been known for their antimicrobial effects indicating the role of RNases in immune defence. In the light of above, features and therapeutic applications of the typical RNases have been summerized in this review for the attention of healthcare research.
 
    How to Cite:
Gupta Shruti, Singh Sukhdev and Kanwar Shamsher Singh , 2016. An Overview on Ribonucleases and their Therapeutic Effects. Insight Medicine, 1: 1-11
DOI: 10.5567/IMAD-IK.2016.1.11
 


INTRODUCTION
Ribonuclease (RNase) is an omnipotent nuclease which catalyzes the degradation of ribonucleic acid (RNA) into smaller components and it has been widely acknowledged as a therapeutic candidate. RNases play pivotal role in RNA metabolism and regulation of fundamental genes expression in living organisms1. They participate in several cellular functions ranging from DNA replication to protein function and defence against foreign microorganisms. RNase-controlled RNA degradation is a determining step in gene regulation, maturation and turnover which is further associated with progression of cancers and infectious diseases. Cytotoxic effects of RNases are the result of catalytic cleavage of available RNAs, byproducts and non-catalytic electrostatic interactions of exogenous enzymes with cell components.

Broadly, RNases are classified as endo-and exo-ribonucleases. Endo-ribonucleases cleave RNA molecule endo-ribonucleolytically (in 5’-3’ direction) while exo-ribonucleases degrade RNA molecule in 3’-5’ direction2. RNases display antitumor, antiviral, antifungal and immunosuppressive properties3,4. They produce genetic damage in cancer cells and destroy their RNA5. Resulted damaged molecular patterns stimulate immune sensors such as toll-like receptors (TLRs) and activated TLRs provoke immunokines which further induce production of cytokines, growth factors and angiogenic modulators and these determine tumor progression6. The anticancer function of immunotoxins which target normal cells also can be improved by introduction of RNases7. An intervention based on combination of RNase with other anticancer molecules can be a promising therapeutic preparation for effective tumor killing.

Since, the discovery of first RNases in 1961, several types of RNases have been explored till date8. Many of them have been studied in great detail with major emphasis on their applications as therapeutic molecule and RNA sequence determination. Exo-ribonucleases remove nucleotides from the 3’-5’ ends by cleaving the phosphodiester bonds at the ends of the polynucleotide chain. These enzymes are highly specific in their cleavage and produce staggered or blunt ends. Endo-ribonucleases cleave the phosphodiester bonds within the single stranded or double stranded RNA molecules. Features of some well known exo-and endo-ribonucleases are described in the following paragraphs.


EXO-RIBONUCLEASES
RNase PH: RNase PH is a 25 kDa E. coli RNase encoded by the rph gene which maps at 81.7 min at the genome. It was first identified due to its phosphorolytic activity against tRNAs9,10. It requires a divalent cation and uses phosphate as a co-substrate to degrade RNA thereby producing 5’ diphosphates. This RNase displays sequence and functional similarity with other E. coli RNases and poly nucleotide phosphorylase (PNPase) also. The RNase PH plays pivotal role in degradation of structural RNAs and provides a potential explanation for the growth defects caused by the absence of the phosphorolytic RNases11.

Polynucleotide phosphorylase: PNPase is encoded by the pnp gene and maps at 69 min at the E. coli12. It is associated with the RNase E and degradosome in the cytoplasm13. Like RNase PH, PNPase also uses phosphate as co-substrate to carry out the phosphorolytic cleavage of RNA and nucleotide diphosphate14. PNPase catalyzes the typical 3’-5’ phosphorolysis of RNA and generates nucleoside diphosphate products15,16. The E. coli PNPase activity is blocked by RNA secondary structure while, the Bacillus subtilis PNPase is hindered by RNA hairpin structures. The E. coli PNPase also repairs the 3’ terminal CCA sequence of tRNA which is also executed by tRNAs17. Crystallographic structure analysis revealed that PNPase is a homotrimeric circular-shaped complex18. Amino acid sequences of bacterial PNPases share high degree of similarity with the PNPases of nuclear genome of plants and mammals19.

RNase II: The RNase II family exo-ribonucleases are present in all domains and degrade RNA from the 3’-end releasing 5’-nucleotide monophosphates20 They participate in the processing, degradation and quality control of all types of RNAs. The E. coli RNase II (72 kDa) is a prototype member of RNase II family exoribonucleases. It is encoded by the rnb gene which is mapped at 29 min on chromosome21. The RNase II has several orthologs and Saccharomyces cerevisiae RNase Rrp44p is one of them. Structural analyses revealed that RNase II contains two N-terminal Cold Shock Domains (CSDs), one C-terminal S1 domain and a central catalytic RNB domain21. The RNase II is a Mg2+ dependent enzyme and its activity is also inhibited by the RNA secondary structure22. The RNase II expression is regulated at transcriptional and post-transcriptional levels23. The RNase II is essential for growth as mutations in RNase II genes have been demonstrated with abnormal chloroplast biogenesis, mitotic control and cancer24,25.

RNase R: It is a 92 kDa RNase encoded by the rnr gene which maps at 95 min in the E. coli genome26,27. The RNase R degrades linear and Y-structure RNAs and doesn’t act on the loop portion of a lariat RNAs. RNase R exhibits 60% sequence homology with RNase II28. It has two N-terminal Cold Shock Domains (CSDs), a central nuclease domain and an S1 domain near the C-terminus29 nuclease domain executes nucleotide degradation wheras CSD and S1 domains give stability to the catalysis. The RNase R expression is essential for the virulence of Shigella sp. and E. coli strains30.

RNase D: RNase D (49 kDa) is encoded by the rnd gene mapped at 40 min on the E. coli chromosome31. It belongs to the DEDD superfamily RNases and performs both DNA and RNA degrading functions32. The RNase D is a divalent metal ion (e.g., Mg2+, Mn2+ and Co2+) dependent RNase and generate ribonucleoside 5’-monophosphate products. The RNase D plays significant roles in tRNA and 5S rRNAs processing also33. The RNase D contains a catalytic domain and two helical domains which come together and form a ring shaped structure33.

RNase T: RNase T is a 23.5 kDa enzyme encoded by the rnt gene (maps at 36 min) in E. coli34. It displays tremendous ribonucleolytic activity among the discovered exo-ribonucleases. It also is a member of DEDD superfamily which is a large family of 3’-5’ exo-nucleases32. It is made up of opposing dimers and functions on tRNA to yield mature 3’end of 5S and 23S rRNA35.

RNase BN and oligo RNase: The RNase BN (encoded by rbn gene) is a 60 kDa ribonuclease which performs 3’ end maturation of tRNAs36. Oligoribonuclease acts on small oligonucleotides and is encoded by the orn gene which maps at 94 min on the E. coli chromosome37. It is a K2 dimer and essential to complete the degradation process of mRNA38.


ENDO-RIBONUCLEASES
RNase I and III: The rna gene at 4.3 min on the chromosome of E. coli encodes a 27 kDa RNase I39 which cleaves within unstructured regions of RNA and forms 2’-3’ cyclic phosphodiester RNA termini. RNase I plays crucial role in the turnover of RNAs and does not dependent on the divalent cations for the hydrolysis function40. The RNase III is an essential enzyme for RNA processing and post-transcriptional gene regulation41. It is a Mg2+-dependent nuclease and is encoded by the rnc gene which maps at 55 min on the chromosome in E. coli. This RNase cleaves phosphodiester bonds of double stranded (ds) RNA and generates 3’ hydroxyl and 5’ phosphate termini. It is a 52 kDa homodimer RNase and contains an N-terminal nuclease domain and a C-terminal dsRNA binding domain (dsRBD). Several orthologs of RNase III have been discovered in prokaryotes and eukaryotes42. The eukaryotic ortholog dicer process the dsRNAs in short interfering (si) RNAs which target RNAs having complementary sequences.

RNase E, P and HI: RNase E is another important RNase for the processing of mRNA, rRNA and tRNA43. It is a 180 kDa Mg2+-dependent phosphodiesterase encoded by rne gene, which maps at 24 min on the chromosome and cleaves adenine and uracil-rich sequences, generating 5’ phosphate and 3’ hydroxyl termini44. Catalytic site of RNase E is present in N-terminal half and C-terminal half contains the RNA binding site45. The RNase P is a divalent cation-dependent ribonucleoprotein RNase and is made up of one RNA subunit and one or more protein subunits46. This ribozyme is encoded by two genes; the protein subunit is coded by rnpA gene (maps at 83 min) and the RNA subunit is coded by rnpB gene which maps at 70 min47.

RNase HI is also a Mg2+-dependent phosphodiesterase that cleaves the RNA strand of RNA-DNA hybrids and is encoded by the rnhA gene (maps at 5 min) in E. coli chromosome48,49. The RNase HI plays pivotal role in ColE1 plasmid replication. The rnhB gene maps at 4.5 min and encodes another ribonuclease namely RNase HII50.


THERAPEUTIC APPLICATIONS OF RNases
Anticancer effects: Anticancer effects of RNases have been extensively studied with animal ribonucleolytic enzymes, viz., bovine pancreatic RNase A, bovine seminal RNase (BS-RNase), onconase and angiogenin51. Among these, BS-RNase and onconase demonstrated significant anticancer potential with bovine pancreatic RNase A, a modest angiogenin activity was observed to work in opposite direction and initiate vascularization of tumor and subsequent tumor growth52. Bovine seminal plasma RNase, onconase and binase are well known anticancer RNases which have been described in following paragraphs. The major types of RNase with their therapeutic applications have been illustrated in Table 1. RNases have the capacity to degrade mRNAs efficiently and thus prevent their translation into biologically active proteins (Fig. 1).

Bovine seminal ribonuclease: The BS-RNase (EC 3.1.27.5) is expressed in the seminal vesicles and testes of Bos taurus53. It is a secretory ribonuclease54. Its native form exists as a dimer in which both subunits are held together by two disulfide linkages between Cys-31 and Cys-3255.

Table 1: Major types of RNases, their characteristics and applications

Figure 1: Action of RNase on the RNA in the target cell

The disulfide linkage undergoes cleavage under reducing conditions of cytosol and the dimer is converted into two monomers. The monomeric form is neutralized by the ribonuclease inhibitor56. Amino acid sequence and crystallographic analysis revealed that BS-RNase belongs to the pancreatic RNase A superfamily57. Each subunit of BS-RNase shares 82% sequence identity with RNase A58. The BS-RNase exerts antiproliferative and apoptotic effects on cancer cells via Beclin-1-mediated autophagy54. It has cytotoxic, aspermagenic and immunosuppressive properties for protecting sperm cells from the female immune system. The BS-RNase suppresses the activation of proliferating lymphocytes by reducing the expression of interleukin (IL-2)59. The BS-RNase treated lymphocytes undergo apoptosis via DNA fragmentation, chromatin migration, disorganised mitochondria and cell shrinking60. The BS-RNase induces cell death in thyroid carcinoma cells also61.

Onconase: Onconase is a 104 amino acid member of RNase A superfamily. It is a promising candidate for the treatment of malignant mesothelioma62. It was isolated from oocytes and early embryos of Rana pipiens. It shares ~30% homology with RNase A and resembles in 3D structure also. Crystallographic and homology studies revealed that onconase has three disulfide linkages at positions 19-68, 30-75 and 48-90 and an imidazol ring of His97 residue rotated at 180° angle63,51. Catalytic site of onconase composed of His10, Lys31 and His97 residues64. Onconase tolerates thermal and guanidine induced transitions (up to 90̊C and 4.4 M) and does not interact with mammalian RNase inhibitor hence evade the attack of inhibitor65. Onconase demonstrated significant cytotoxic effects against cancer cell lines i.e., HL-60, HT-29, 9L rat glioma, K-562, Colo-320, JCA-1, U937, A549 and ASPC-166,67. Onconase and its derived products exhibited potent antitumor effects against cervical, breast, colon, pancreatic, ovarian and prostate cancers with LD50 (median lethal dose) value ~10–7 M3. It catalyzes the formation of interfering RNAs (RNAi), degrades tRNAs and inhibits protein synthesis, which results in apoptosis of the cell68. Onconase modulates cytokine-receptor interactions, MAPK, Jak-STAT, Bcl-2, Bax and various other signaling pathways in cancer models69. It activates jun-N-terminal kinase (JNK) and caspase-9, -3 and -7 proteins in HeLa cells, serine proteases in HL-60 cells64 and IL-6, IL-24 and ATF-3 in MM cell line69. In lymphocytic leukemia onconase mediates its apoptotic effects by reducing NF-κB expression level70. Onconase also pull down the level of reactive oxygen species in cancer cells71. The cytotoxic potential of onconase increased when positively charged residues were added to the enzyme by site-directed mutagenesis and chemical modification72,73. Onconase modulates tumor cell apoptosis at microRNA expression level and reduces the oncogenic microRNAs in malignant mesothelioma models74,65. In clinical trials onconase demonstrated some adverse effects, viz., lymphocyte proliferation suppression, renal failure, bone marrow toxicity, mascular stiffness and tremor75.

Binase: Bacteria also provide anticancer RNases3,83. Several bacterial species producing RNase with cytotoxicity towards cancer models have been discovered and Bacillus intermedius84, Bacillus amyloliquefaciens85,86 and Streptomyces aureofaciens87 are exemplary among them. Binase (EC 3.1.27.3) is a 12.2 kDa extracellular cationic RNase from Bacillus intermedius77. It cleaves RNA at purine residues and does not require any cofactor to accomplish this hydrolysis86,87. Binase was the first anticancer bacterial RNase which demonstrated comparable cytotoxic potential against malignant cells62. Binase exhibited significant antiproliferative and apoptotic activities on K562, A549 and ovary cancer cells77. It elicited cell death in transformed fibroblasts and myeloid progenitor cells88 and demonstrated cytotoxic effects on Kasumi-1 cells with half-maximal concentration of 0.56 μM. Binase has suppressive effects on several oncogenes also i.e., KIT, AML1-ETO and FLT3-ITD89. Moreover, Binase has low immunogenicity90 and doesn’t affect cell viability of leukocytes and myeloid progenitor cells91. Binase also demonstrated antiviral properties against rabies viruses, plant viruses and the influenza strains78.

Some other RNases have also been reported with anticancer activities. RNase L showed antiproliferative effects against H9 leukemia cells92. RNase Sa3 from Streptomyces aureofaciens93 showed cytotoxicity against K562 cells with IC50 of 5 μM. RNase Sa3 is not inhibited by the cytosolic RNase inhibitor94. Some of the mushroom species also have been known for their anticancer properties. Hypsizigus marmoreus RNase (18 kDa) reduced the L1210 proliferation (IC50 60 μM). Another RNase (14.5 kDa) from fresh fruiting bodies of the edible mushroom Lyophyllum shimeji exhibited cytostatic potential on liver cancer HepG2 cells (IC50 10 μM) and on breast cancer MCF7 cells (IC50 6.2 μM)95. A 28 kDa RNase from ascocarps of Tuber indicum showed antiproliferative effects on HepG2 and MCF7 cells with IC50 values 12.6 and 16.6 μM, respectively96.


RNases IN HOST DEFENCE
Eosinophil RNases: Cytoplasmic granules of human eosinophilic leukocytes secrete two major ribonuclease proteins97, the Eosinophil Cationic Protein (ECP) and the Eosinophil Derived Neurotoxin (EDN). The ECP and EDN belong to the pancreatic type RNase family and share 70 and 90% similarity in their amino acid and nucleotide sequences98. The EDN is also known as RNase II or eosinophilic protein-X which demonstrated better RNase activity than ECP99,100. The EDN is located on ‘q’ arm of chromosome 14101 and shares similarity with human liver and urinary RNase (RNase U)102. The EDN is an 18.6 kDa single chain polypeptide with four characteristic disulfide bonds and His15-Lys38-His129 catalytic triad103,104. The RNases can elicit immune response via leukocyte activation, maturation and chemotaxis. The EDN showed antiviral activities against respiratory syncytial virus, HIV-1 and some RNA viruses81,82. The recombinant EDN (EDNsFv) created by fusing human EDN gene and antibody fragment of human transferring receptor exhibited significant cytotoxic effects against transferring receptor expressing leukemia cells105. Another recombinant EDN (rhEDN) reduced the infectivity of respiratory syncytial virus which causes asthma aggravations106,107. The EDN levels correlate with neuroinflammation characteristic in Amyotrophic Lateral Sclerosis (ALS), a neurodegenerative disorder. Thus EDN is used as a biomarker for ALS disease81. The EDN engineered with hepatitis B virus core protein (HBVc), suppressed the hepatitis B infected cells without affecting normal cells RNA105. Some other ribonucleases i.e., mEar 11 and mEar 2 (the mouse eosinophil associated RNases) are also reported for antiviral and immunogenic effects. Alveolar macrophages produce mEar 11 upon administration of IL-4 or 13. The mEar 11 is a chemo-attractant for CD11c+ dendritic cells and F4/80+CD11c- macrophages108. These mEars exhibited significant antiviral effects against influenza strains and pneumonia virus of mice109.

RNase 7 and 8: RNase 7 (~14.5 kDa) is a member of RNase A superfamily and the gene encoding RNase 7 is located on chromosome110 14q11.2. It is expressed in skin, liver, kidney, skeletal muscles and heart99. It exhibited potential antibacterial activities against Enterococcus faecium, Pseudomonas aeruginosa and Pichia pastoris79. Cationic residues in RNase 7 bind to negatively charged components on bacterial surface and facilitate the insertion of RNase. In P. aeruginosa, RNase 7 enters the cell by making complex with the outer membrane protein Opr I111. The RNase 7 was reported to be express significantly in response to the external stimuli112. Wanke and colleagues113 studied the host defence aspects of RNase 7 that Staphylococcus epidermis induces RNase 7 expression in keratocytes via TLR2, EGFR and NF-κB pathways. Studies with protozoan’s exposure also demonstrated that RNase 7 contributes to a responsible role in host defence114. RNase 8 is another ribonuclease in the RNase A superfamily. It shares 78% amino acid similarity with RNase 7115. The RNase 8 play responsible role in placental host defence by defending the foetus from pathogen from the maternal circulation116. RNase 8 exhibited significant antimicrobial activities against Klebsiella pneumonia, Enterococcus faecium, E. faecalis, Staphylococcus aureus, Pseudomonas aeruginosa and Candida albicans117.

Angiogenin: Angiogenin is a 14 kDa ribonuclease which was isolated from HT-29 conditioned media116,118. It has the ability to induce the formation of new blood vessels119,120. Angiogenin has a receptor binding site (which facilitates the angiogenin entry in the cell), a nuclear localization sequence (by which angiogenin enters the nucleus) and a catalytic site which catalyze the tRNA cleavage121. Angiogenin is reported for its host defence features. In mouse, six kinds of angiogenin (1-6) are reported. Mouse angiogenin 4 is reported to be significantly expressed in paneth cells upon bacterial LPS (lipopolysaccharide) challenge. It exhibited significant antibacterial activities against intestinal microbes80,99.

Besides these applications, RNases have been reported to exhibit antiviral functions also. RNase from Rana catesbeiana suppressed the multiplication of Japanese encephalitis virus and accelerated apoptosis of virus-infected cells122. Yadav and Batra123 recognized specific targets of restrictocin, an RNase from Aspergillus restrictus in HIV-1 genome. Some mushroom RNases displayed inhibitory effects against Reverse Transcriptase (RT) of HIV-13. The RNases from mushroom species Thelephora ganbajun (~30 kDa)124, Lyophyllum shimehi96, Hygrophorus russula (~28 kDa)125, Hohenbuehelia serotina (~27 kDa)4 and Ramaria formosa (~29 kDa)126 inhibited HIV-1-RT with IC50 concentrations 0.3, 7.2, 4.64, 50 and 3 μM, respectively.


CONCLUSION
Ribonucleases are potential therapeutic candidates and must be converted into druggable forms with sufficient bioavailability. They have promising anticancer and antiviral applications. Till date, very low number of bacterial RNases have been discovered. Also, none of the discovered RNase has crossed the clinical barriers for therapeutic use. Genetic pathways of their synthesis and mechanisms of actions in cancer cells are still a topic of research which needs to be understood for their development into a therapeutic product. So, there is an essential need to discover potential and elaborate the ribonucleases of therapeutic value. RNases mentioned in this review must be further researched for clinical trials and druggability evaluation especially BS-RNase and onconase for anticancer value and RNase 7, 8 and eosinophil RNases for anti-HIV properties. In final words, this review provides general information about RNases and can help the healthcare pharmacy for the treatment of serious metabolic syndromes including cancers and AIDS.


ACKNOWLEDGMENTS
Financial support from Department of Biotechnology, Ministry of Science and Technology, Govt. of India to Mr. Sukhdev Singh (Grant No. DBT-JRF/F-19/487) is gratefully acknowledged.


REFERENCES

  1. Kocic, G., G. Bjelakovic, D. Pavlovic, T. Jevtovic and V. Pavlovic et al., 2007. Protective effect of interferon-α on the DNA- and RNA-degrading pathway in anti-Fas-antibody induced apoptosis. Hepatol. Res., 37: 637-646

  2. Lawal, A., O. Jejelowo, A.K. Chopra and J.A. Rosenzweig, 2011. Ribonucleases and bacterial virulence. Microb. Biotechnol., 4: 558-571

  3. Fang, E.F. and T.B. Ng, 2011. Ribonucleases of different origins with a wide spectrum of medicinal applications. Biochimica Biophysica Acta (BBA)-Rev. Cancer, 1815: 65-74

  4. Zhang, R., L. Zhao, H. Wang and T.B. Ng, 2014. A novel ribonuclease with antiproliferative activity toward leukemia and lymphoma cells and HIV-1 reverse transcriptase inhibitory activity from the mushroom, Hohenbuehelia serotina. Int. J. Mol. Med., 33: 209-214

  5. Huang, Y.C., Y.M. Lin, T.W. Chang, S.J. Wu and Y.S. Lee et al., 2007. The flexible and clustered lysine residues of human ribonuclease 7 are critical for membrane permeability and antimicrobial activity. J. Biol. Chem., 282: 4626-4633

  6. Mitkevich, V.A., O.N. Ilinskaya and A.A. Makarov, 2015. Antitumor RNases: Killer’s secrets. Cell Cycle, 14: 931-932

  7. Schirrmann, T., J. Krauss, M.A.E. Arndt, S.M. Rybak and S. Dubel, 2009. Targeted therapeutic RNases (ImmunoRNases). Expert Opin. Biol. Ther., 9: 79-95

  8. Spahr, P.F. and B.R. Hollingworth, 1961. Purification and mechanism of action of ribonuclease from Escherichia coli Ribosomes. J. Biol. Chem., 236: 823-831

  9. Li, Z. and M.P. Deutscher, 1996. Maturation pathways for E. coli tRNA precursors: A random multienzyme process in vivo. Cell, 86: 503-512

  10. Leszczyniecka, M., R. DeSalle, D.C. Kang and P.B. Fisher, 2004. The origin of polynucleotide phosphorylase domains. Mol. Phylogenet. Evol., 31: 123-130

  11. Jain, C., 2012. Novel role for RNase PH in the degradation of structured RNA. J. Bacteriol., 194: 3883-3890

  12. Houseley, J., J. LaCava and D. Tollervey, 2006. RNA-quality control by the exosome. Nat. Rev. Mol. Cell. Biol., 7: 529-539

  13. Marcaida, M.J., M.A. DePristo, V. Chandran, A.J. Carpousis and B.F. Luisi, 2006. The RNA degradosome: Life in the fast lane of adaptive molecular evolution. Trends Biochem. Sci., 31: 359-365

  14. Portnoy, V. and G. Schuster, 2006. RNA polyadenylation and degradation in different Archaea; roles of the exosome and RNase R. Nucl. Acids Res., 34: 5923-5931

  15. Buttner, K., K. Wenig and K.P. Hopfner, 2006. The exosome: A macromolecular cage for controlled RNA degradation. Mol. Microbiol., 61: 1372-1379

  16. Sarkar, D. and P.B. Fisher, 2006. Polynucleotide phosphorylase: An evolutionary conserved gene with an expanding repertoire of functions. Pharmacol. Ther., 112: 243-263

  17. Lin-Chao, S., N.T. Chiou and G. Schuster, 2007. The PNPase, exosome and RNA helicases as the building components of evolutionarily-conserved RNA degradation machines. J. Biomed. Sci., 14: 523-532

  18. Shi, Z., W.Z. Yang, S. Lin-Chao, K.F. Chak and H.S. Yuan, 2008. Crystal structure of Escherichia coli PNPase: Central channel residues are involved in processive RNA degradation. RNA. Soc., 14: 2361-2371

  19. Portnoy, V., G. Palnizky, S. Yehudai-Resheff, F. Glaser and G. Schuster, 2008. Analysis of the human polynucleotide phosphorylase (PNPase) reveals differences in RNA binding and response to phosphate compared to its bacterial and chloroplast counterparts. RNA., 14: 297-309

  20. Grossman, D. and A. van Hoof, 2006. RNase II structure completes group portrait of 3’ exoribonucleases. Nat. Struct. Mol. Biol., 13: 760-761

  21. Arraiano, C.M., R.G. Matos and A. Barbas, 2010. RNase II: The finer details of the Modus operandi of a molecular killer. RNA. Biol., 7: 276-281

  22. LaCava, J., J. Houseley, C. Saveanu, E. Petfalski, E. Thompson, A. Jacquier and D. Tollervey, 2005. RNA degradation by the exosome is promoted by a nuclear polyadenylation complex. Cell, 121: 713-724

  23. Schneider, C., E. Leung, J. Brown and D. Tollervey, 2009. The N-terminal PIN domain of the exosome subunit Rrp44 harbors endonuclease activity and tethers Rrp44 to the yeast core exosome. Nucl. Acids Res., 37: 1127-1140

  24. Bollenbach, T.J., H. Lange, R. Gutierrez, M. Erhardt, D.B. Stern and D. Gagliardi, 2005. RNR1, a 3’ 5’ exoribonuclease belonging to the RNR superfamily, catalyzes 3’ maturation of chloroplast ribosomal RNAs in Arabidopsis thaliana. Nucl. Acids Res., 33: 2751-2763

  25. Andrade, J.M., E. Hajnsdorf, P. Regnier and C.M. Arraiano, 2009. The poly(A)-dependent degradation pathway of rpsO mRNA is primarily mediated by RNase R. RNA., 15: 316-326

  26. Cheng, Z.F., Y. Zuo, Z. Li, K.E. Rudd and M.P. Deutscher, 1998. The vacB gene required for virulence in Shigella flexneri and Escherichia coli encodes the exoribonuclease RNase R. J. Biol. Chem., 273: 14077-14080

  27. Li, Z., S. Reimers, S. Pandit and M.P. Deutscher, 2002. RNA quality control: Degradation of defective transfer RNA. EMBO J., 21: 1132-1138

  28. Cheng, Z.F. and M.P. Deutscher, 2003. Quality control of ribosomal RNA mediated by polynucleotide phosphorylase and RNase R. Proc. Natl. Acad. Sci. USA., 100: 6388-6393

  29. Suzuki, H., Y. Zuo, J. Wang, M.Q. Zhang, A. Malhotra and A. Mayeda, 2006. Characterization of RNase R-digested cellular RNA source that consists of lariat and circular RNAs from pre-mRNA splicing. Nucl. Acids Res., Vol. 34, No. 8. 10.1093/nar/gkl151

  30. Vincent, H.A. and M.P. Deutscher, 2009. Insights into how RNase R degrades structured RNA: Analysis of the nuclease domain. J. Mol. Biol., 387: 570-583

  31. Zhang, J. and M.P. Deutscher, 1988. Escherichia coli RNase D: Sequencing of the rnd structural gene and purification of the overexpressed protein. Nucl. Acids Res., 16: 6265-6278

  32. Zuo, Y. and M.P. Deutscher, 2001. Exoribonuclease superfamilies: Structural analysis and phylogenetic distribution. Nucl. Acids Res., 29: 1017-1026

  33. Zuo, Y., Y. Wang and A. Malhotra, 2005. Crystal structure of Escherichia coli RNase D, an exoribonuclease involved in structured RNA processing. Structure, 13: 973-984

  34. Li, Z., S. Pandit and M.P. Deutscher, 1998. 3′ Exoribonucleolytic trimming is a common feature of the maturation of small, stable RNAs in Escherichia coli. Proc. Natl. Acad. Sci. USA., 95: 2856-2861

  35. Zuo, Y., H. Zheng, Y. Wang, M. Chruszcz and M. Cymborowski et al., 2007. Crystal structure of RNase T, an exoribonuclease involved in tRNA maturation and end turnover. Structure, 15: 417-428

  36. Schilling, O., S. Ruggeberg, A. Vogel, N. Rittner and S. Weichert et al., 2004. Characterization of an Escherichia coli elaC deletion mutant. Biochem. Biophys. Res. Commun., 320: 1365-1373

  37. Mechold, U., V. Ogryzko, S. Ngo and A. Danchin, 2006. Oligoribonuclease is a common downstream target of lithium-induced pAp accumulation in Escherichia coli and human cells. Nucl. Acid Res., 34: 2364-2373

  38. Dutta, T., A. Malhotra and M.P. Deutscher, 2012. Exoribonuclease and endoribonuclease activities of RNase BN/RNase Z both function in vivo. J. Biol. Chem., 287: 35747-35755

  39. Meador, III J. and D. Kennell, 1990. Cloning and sequencing the gene encoding Escherichia coli ribonuclease I: Exact physical mapping using the genome library. Gene, 95: 1-7

  40. Cannistraro, V.J. and D. Kennell, 1991. RNase I*, a form of RNase I and mRNA degradation in Escherichia coli. J. Bacteriol., 173: 4653-4659

  41. Lamontagne, B., S. Larose, J. Boulanger and S.A. Elela, 2001. The RNase III family: A conserved structure and expanding functions in eukaryotic dsRNA metabolism. Curr. Issues Mol. Biol., 3: 71-78

  42. Xiao, J., C.E. Feehery, G. Tzertzinis and C.V. Maina, 2009. E. coli RNase III(E38A) generates discrete-sized products from long dsRNA. RNA, 15: 984-991

  43. Clarke, J.E., L. Kime, A.D. Romero and K.J. McDowall, 2014. Direct entry by RNase E is a major pathway for the degradation and processing of RNA in Escherichia coli. Nucl. Acids Res., 42: 11733-11751

  44. Kaberdin, V.A., 2003. Probing the substrate specificity of Escherichia coli RNase E using a novel oligonucleotide-based assay. Nucl. Acids Res., 31: 4710-4716

  45. Kime, L., J.E. Clarke, A.D. Romero, J.A. Grasby and K.J. McDowall, 2014. Adjacent single-stranded regions mediate processing of tRNA precursors by RNase E direct entry. Nucl. Acids Res., 42: 4577-4589

  46. Ando, T., T. Tanaka and Y. Kikuchi, 2003. Substrate shape specificity of E. coli RNase P ribozyme is dependent on the concentration of magnesium ion. J. Biochem., 133: 445-451

  47. Gopalan, V., A. Vioque and S. Altman, 2002. RNase P: Variations and uses. J. Biol. Chem., 277: 6759-6762

  48. Mian, I.S., 1997. Comparative sequence analysis of ribonucleases HII, III, II, PH and D. Nucl. Acids Res., 25: 3187-3195

  49. Nicholson, A.W., 1997. Escherichia coli Ribonucleases: Paradigms for Understanding Cellular RNA Metabolism and Regulation. In: Ribonucleases: Structures and Function, D’Alessio, G. and J.F. Riordan (Eds.). Academic Press, New York, USA., ISBN: 9780080540597, pp: 1-49.

  50. Haruki, M., K. Hayashi, T. Kochi, A. Muroya and Y. Koga et al., 1998. Gene cloning and characterization of recombinant RNase HII from a hyperthermophilic archaeon. J. Bacteriol., 180: 6207-6214

  51. Ardelt, W., B. Ardelt and Z. Darzynkiewicz, 2009. Ribonucleases as potential modalities in anticancer therapy. Eur. J. Pharmacol., 625: 181-189

  52. Matousek, J. and J. Matousek, 2010. Plant ribonucleases and nucleases as antiproliferative agens targeting human tumors growing in mice. Rec. Patents DNA Gene Sequences, 4: 29-39

  53. Eller, C.H., J.E. Lomax and R.T. Raines, 2014. Bovine brain ribonuclease is the functional homolog of human ribonuclease 1. J. Biol. Chem., 289: 25996-26006

  54. Fiorini, C., G. Gotte, F. Donnarumma, D. Picone and M. Donadelli, 2014. Bovine seminal ribonuclease triggers Beclin1-mediated autophagic cell death in pancreatic cancer cells. Biochimica Biophysica Acta (BBA)-Mol. Cell Res., 1843: 976-984

  55. Gotte, G., A.M. Helmy, C. Ercole, R. Spadaccini, D.V. Laurents, M. Donadelli and D. Picone, 2012. Double domain swapping in bovine seminal RNase: Formation of distinct N-and C-swapped tetramers and multimers with increasing biological activities. PloS One, Vol. 7. 10.1371/journal.pone.0046804

  56. Giancola, C., C. Ercole, I. Fotticchia, R. Spadaccini, E. Pizzo, G. D’Alessio and D. Picone, 2011. Structure-cytotoxicity relationships in bovine seminal ribonuclease: New insights from heat and chemical denaturation studies on variants. FEBS J., 278: 111-122

  57. Dyer, K.D. and H.F. Rosenberg, 2006. The RNase a superfamily: Generation of diversity and innate host defense. Mol. Diversity, 10: 585-597

  58. Spadaccini, R., C. Ercole, M.A. Gentile, D. Sanfelice and R. Boelens et al., 2012. NMR studies on structure and dynamics of the monomeric derivative of BS-RNase: New insights for 3D domain swapping. PloS One, Vol. 7. 10.1371/journal.pone.0029076

  59. Tamburrini, M., G. Scala, C. Verde, M.R. Ruocco, A. Parente, S. Venuta and G. D’Alessio, 1990. Immunosuppressive activity of bovine seminal RNase on T-cell proliferation. Eur. J. Biochem., 190: 145-148

  60. Sinatra, F., D. Callari, M. Viola, M.T. Longombardo and M. Patania et al., 2000. Bovine seminal RNase induces apoptosis in normal proliferating lymphocytes. Int. J. Clin. Lab. Res., 30: 191-196

  61. Spalletti-Cernia, D., R. Sorrentino, S. Di Gaetano, A. Arciello and C. Garbi et al., 2003. Antineoplastic ribonucleases selectively kill thyroid carcinoma cells via caspase-mediated induction of apoptosis. J. Clin. Endocrinol. Metab., 88: 2900-2907

  62. Sen’kova, A.V., N.L. Mironova, O.A. Patutina, V.A. Mitkevich and O.V. Markov et al., 2014. Ribonuclease binase decreases destructive changes of the liver and restores its regeneration potential in mouse lung carcinoma model. Biochimie, 101: 256-259

  63. Ardelt, W., K. Shogen and Z. Darzynkiewicz, 2008. Onconase and amphinase, the antitumor ribonucleases from Rana pipiens oocytes. Curr. Pharm. Biotechnol., 9: 215-225

  64. Lee, J.E. and R.T. Raines, 2008. Ribonucleases as novel chemotherapeutics: The ranpirnase example. BioDrugs, 22: 53-58

  65. Qiao, M., L.D. Zu, X.H. He, R.L. Shen, Q.C. Wang and M.F. Liu, 2012. Onconase downregulates microRNA expression through targeting microRNA precursors. Cell Res., 22: 1199-1202

  66. Majchrzak, A., M. Witkowska, A. Medra, M. Zwolinska and J. Bogusz et al., 2013. In vitro cytotoxicity of ranpirnase (onconase) in combination with components of R-CHOP regimen against Diffuse Large B Cell Lymphoma (DLBCL) cell line. Postepy Higieny Medycyny Doswiadczalnej, 67: 1166-1172

  67. Wang, X. and Z. Guo, 2015. Chlorotoxin-conjugated onconase as a potential anti-glioma drug. Oncol. Lett., 9: 1337-1342

  68. Halicka, H.D., B. Ardelt, K. Shogen and Z. Darzynkiewicz, 2007. Mild hyperthermia predisposes tumor cells to undergo apoptosis upon treatment with onconase. Int. J. Oncol., 30: 841-847

  69. Altomare, D.A., S.M. Rybak, J. Pei, J.V. Maizel, M. Cheung, J.R. Testa and K. Shogen, 2010. Onconase responsive genes in human mesothelioma cells: Implications for an RNA damaging therapeutic agent. BMC Cancer, Vol. 10. 10.1186/1471-2407-10-34

  70. Ita, M., H.D. Halicka, T. Tanaka, A. Kurose, B. Ardelt, K. Shogen and Z. Darzynkiewicz, 2008. Remarkable enhancement of cytotoxicity of onconase and cepharanthine when used in combination on various tumor cell lines. Cancer Biol. Therapy, 7: 1104-1108

  71. Ardelt, B., W. Ardelt, P. Pozarowski, J. Kunicki, K. Shogen and Z. Darzynkiewicz, 2007. Cytostatic and cytotoxic properties of Amphinase: A novel cytotoxic ribonuclease from Rana pipiens oocytes. Cell Cycle, 6: 3097-3102

  72. Futami, J. and H. Yamada, 2008. Design of cytotoxic ribonucleases by cationization to enhance intracellular protein delivery. Curr. Pharm. Biotechnol., 9: 180-184

  73. Turcotte, R.F., L.D. Lavis and R.T. Raines, 2009. Onconase cytotoxicity relies on the distribution of its positive charge. FEBS J., 276: 3846-3857

  74. Goparaju, C.M., J.D. Blasberg, S. Volinia, J. Palatini and S. Ivanov et al., 2011. Onconase mediated NFKβ downregulation in malignant pleural mesothelioma. Oncogene, 30: 2767-2777

  75. Weber, T., A. Mavratzas, S. Kiesgen, S. Haase and B. Botticher et al., 2015. A humanized anti-CD22-onconase antibody-drug conjugate mediates highly potent destruction of targeted tumor cells. J. Immunol. Res., Vol. 2015. 10.1155/2015/561814

  76. Zwolinska, M. and P. Smolewski, 2010. [Onconase: A ribonuclease with antitumor activity]. Postepy Higieny Medycyny Doswiadczalnej, 64: 58-66, (In Polish)

  77. Garipov, A.R., A.A. Nesmelov, H.A. Cabrera-Fuentes and O.N. Ilinskaya, 2014. Bacillus intermedius ribonuclease (BINASE) induces apoptosis in human ovarian cancer cells. Toxicon, 92: 54-59

  78. Mahmud, R.S. and O.N. Ilinskaya, 2013. Antiviral activity of binase against the pandemic influenza A (H1N1) virus. Acta Naturae, 5: 44-51

  79. Spencer, J.D., A.L. Schwaderer, H. Wang, J. Bartz and J. Kline et al., 2013. Ribonuclease 7, an antimicrobial peptide upregulated during infection, contributes to microbial defense of the human urinary tract. Kidney Int., 83: 615-625

  80. Li, S. and G.F. Hu, 2010. Angiogenin-mediated rRNA transcription in cancer and neurodegeneration. Int. J. Biochem. Mol. Biol., 1: 26-35

  81. Liu, G.T., C.S. Hwang, C.H. Hsieh, C.H. Lu and S.L.Y. Chang et al., 2013. Eosinophil-derived neurotoxin is elevated in patients with amyotrophic lateral sclerosis. Med. Inflamm. 10.1155/2013/421389

  82. Yang, D., Q. Chen, S.B. Su, P. Zhang and K. Kurosaka et al., 2008. Eosinophil-derived neurotoxin acts as an alarmin to activate the TLR2-MyD88 signal pathway in dendritic cells and enhances Th2 immune responses. J. Exp. Med., 205: 79-90

  83. Mitkevich, V.A., O.V. Kretova, I.Y. Petrushanko, K.M. Burnysheva and D.V. Sosin et al., 2013. Ribonuclease binase apoptotic signature in leukemic Kasumi-1 cells. Biochimie, 95: 1344-1349

  84. Ilinskaya, O.N., A. Koschinski, H. Repp, V.A. Mitkevich and F. Dreyer et al., 2008. RNase-induced apoptosis: Fate of calcium-activated potassium channels. Biochimie, 90: 717-725

  85. Edelweiss, E., T.G. Balandin, J.L. Ivanova, G.V. Lutsenko and O.G. Leonova et al., 2008. Barnase as a new therapeutic agent triggering apoptosis in human cancer cells. PloS One, Vol. 3. 10.1371/journal.pone.0002434

  86. Ulyanova, V., V. Vershinina and O. Ilinskaya, 2011. Barnase and binase: Twins with distinct fates. FEBS J., 278: 3633-3643

  87. Cabrera-Fuentes, H.A., M. Aslam, M. Saffarzadeh, A. Kolpakov, P. Zelenikhin, K.T. Preissner and O.N. Ilinskaya, 2013. Internalization of Bacillus intermedius ribonuclease (BINASE) induces human alveolar adenocarcinoma cell death. Toxicon, 69: 219-226

  88. Mitkevich, V.A., N.A. Tchurikov, P.V. Zelenikhin, I.Y. Petrushanko, A.A. Makarov and O.N. Ilinskaya, 2010. Binase cleaves cellular noncoding RNAs and affects coding mRNAs. FEBS J., 277: 186-196

  89. Mitkevich, V.A., I.Y. Petrushanko, P.V. Spirin, T.V. Fedorova and O.V. Kretova et al., 2011. Sensitivity of acute myeloid leukemia Kasumi-1 cells to binase toxic action depends on the expression of KIT and АML1-ETO oncogenes. Cell Cycle, 10: 4090-4097

  90. Mironova, N., I. Petrushanko, O. Patutina, A.V. Sen’kova and O. Simonenko et al., 2013. Ribonuclease binase inhibits primary tumor growth and metastases via apoptosis induction in tumor cells. Cell Cycle, 12: 2120-2131

  91. Makarov, A.A., A. Kolchinsky and O.N. Ilinskaya, 2008. Binase and other microbial RNases as potential anticancer agents. BioEssays, 30: 781-790

  92. Le Roy, F., C. Bisbal, M. Silhol, C. Martinand, B. Lebleu and T. Salehzada, 2001. The 2-5A/RNase L/RNase L inhibitor (RNI) pathway regulates mitochondrial mRNAs stability in interferon α-treated H9 cells. J. Biol. Chem., 276: 48473-48482

  93. Hlinkova, V., L. Urbanikova, D. Krajcikova and J. Sevcik, 2001. Purification, crystallization and preliminary X-ray analysis of two crystal forms of ribonuclease Sa3. Acta Crystallogr. Sect. D: Biol. Crystallogr., 57: 737-739

  94. Sevcik, J., L. Urbanikova, P.A. Leland and R.T. Raines, 2002. X-ray structure of two crystalline forms of a Streptomycete ribonuclease with cytotoxic activity. J. Biol. Chem., 277: 47325-47330

  95. Zhang, R.Y., G.Q. Zhang, D.D. Hu, H.X. Wang and T.B. Ng, 2010. A novel ribonuclease with antiproliferative activity from fresh fruiting bodies of the edible mushroom Lyophyllum shimeiji. Biochem. Genet., 48: 658-668

  96. Xiao, C., S.S. Feng, H.X. Wang, Z.Y. Gong and T.B. Ng, 2014. Purification and characterization of a ribonuclease with antiproliferative activity from the mystical wild mushroom Tuber indicum. J. Basic Microbiol., 54: S102-S108

  97. Domachowske, J.B., K.D. Dyer, A.G. Adams, T.L. Leto and H.F. Rosenberg, 1998. Eosinophil cationic protein/RNase 3 is another RNase A-family ribonuclease with direct antiviral activity. Nucl. Acids Res., 26: 3358-3363

  98. Navarro, S., J. Aleu, M. Jimenez, E. Boix, C.M. Cuchillo and M.V. Nogues, 2008. The cytotoxicity of eosinophil cationic protein/ribonuclease 3 on eukaryotic cell lines takes place through its aggregation on the cell membrane. Cell. Mol. Life Sci., 65: 324-337

  99. Rosenberg, H.F., 2008. RNase A ribonucleases and host defense: An evolving story. J. Leukoc. Biol., 83: 1079-1087

  100. Fang, S.L., T.C. Fan, H.W. Fu, C.J. Chen and C.S. Hwang et al., 2013. A novel cell-penetrating peptide derived from human eosinophil cationic protein. PLoS One, Vol. 8. 10.1371/journal.pone.0057318

  101. Cho, S., J.J. Beintema and J. Zhang, 2005. The ribonuclease a superfamily of mammals and birds: Identifying new members and tracing evolutionary histories. Genomics, 85: 208-220

  102. Sorrentino, S., D.G. Glitz, K.J. Hamann, D.A. Loegering, J.L. Checkel and G.J. Gleich, 1992. Eosinophil-derived neurotoxin and human liver ribonuclease. Identity of structure and linkage of neurotoxicity to nuclease activity. J. Biol. Chem., 267: 14859-14865

  103. Swaminathan, G.J., D.E. Holloway, K. Veluraja and K.R. Acharya, 2002. Atomic resolution (0.98 A) structure of eosinophil-derived neurotoxin. Biochemistry, 41: 3341-3352

  104. Kephart, G.M., J.A. Alexander, A.S. Arora, Y. Romero, T.C. Smyrk, N.J. Talley and H. Kita, 2010. Marked deposition of eosinophil-derived neurotoxin in adult patients with eosinophilic esophagitis. Am. J. Gastroenterol., 105: 298-307

  105. Rosenberg, H.F., 2015. Eosinophil-derived neurotoxin (EDN/RNase 2) and the mouse eosinophil-associated RNases (mEars): Expanding roles in promoting host defense. Int. J. Mol. Sci., 16: 15442-15455

  106. Jackson, D.J. and S.L. Johnston, 2010. The role of viruses in acute exacerbations of asthma. J. Allergy Clin. Immunol., 125: 1178-1187

  107. Kumar, R.K., P.S. Foster and H.F. Rosenberg, 2014. Respiratory viral infection, epithelial cytokines and innate lymphoid cells in asthma exacerbations. J. Leukoc. Biol., 96: 391-396

  108. Yamada, K.J., T. Barker, K.D. Dyer, T.A. Rice and C.M. Percopo et al., 2015. Eosinophil-associated ribonuclease 11 is a macrophage chemoattractant. J. Biol. Chem., 290: 8863-8875

  109. Gaudreault, E. and J. Gosselin, 2007. Leukotriene B4-mediated release of antimicrobial peptides against cytomegalovirus is BLT1 dependent. Viral Immunol., 20: 407-420

  110. Koten, B., M. Simanski, R. Glaser, R. Podschun, J.M. Schroder and J. Harder, 2009. RNase 7 contributes to the cutaneous defense against Enterococcus faecium. PLoS One, Vol., 4, No. 7. 10.1371/journal.pone.0006424

  111. Lin, Y.M., S.J. Wu, T.W. Chang, C.F. Wang and C.S. Suen et al., 2010. Outer membrane protein I of Pseudomonas aeruginosa is a target of cationic antimicrobial peptide/protein. J. Biol. Chem., 285: 8985-8994

  112. Mohammed, I., A. Yeung, A. Abedin, A. Hopkinson and H.S. Dua, 2010. Signalling pathways involved in ribonuclease-7 expression. Cell Mol. Life Sci., 68: 1941-1952

  113. Wanke, I., H. Steffen, C. Christ, B. Krismer and F. Gotz et al., 2010. Skin commensals amplify the innate immune response to pathogens by activation of distinct signaling pathways. J. Invest Dermatol., 131: 382-390

  114. Otri, A.M., I. Mohammed, A. Abedin, Z. Cao, A. Hopkinson, N. Panjwani and H.S. Dua, 2010. Antimicrobial peptides expression by ocular surface cells in response to Acanthamoeba castellanii: An in vitro study. Br. J. Ophthalmol., 94: 1523-1527

  115. Zhang, J., K.D. Dyer and H.F. Rosenberg, 2002. RNase 8, a novel RNase A superfamily ribonuclease expressed uniquely in placenta. Nucl. Acids Res., 30: 1169-1175

  116. Gupta, S.K., B.J. Haigh, F.J. Griffin and T.T. Wheeler, 2012. The mammalian secreted RNases: Mechanisms of action in host defence. Innate Immun., 19: 86-97

  117. Rudolph, B., R. Podschun, H. Sahly, S. Schubert, J.M. Schroder and J. Harder, 2006. Identification of RNase 8 as a novel human antimicrobial protein. Antimicrob. Agents Chemother., 50: 3194-3196

  118. Fu, H., J. Feng, Q. Liu, F. Sun and Y. Tie et al., 2009. Stress induces tRNA cleavage by angiogenin in mammalian cells. FEBS Lett., 583: 437-442

  119. Yamasaki, S., P. Ivanov, G.F. Hu and P. Anderson, 2009. Angiogenin cleaves tRNA and promotes stress-induced translational repression. J. Cell Biol., 185: 35-42

  120. Dutta, S., C. Bandyopadhyay, V. Bottero, M.V. Veettil and L. Wilson et al., 2014. Angiogenin interacts with the plasminogen activation system at the cell surface of breast cancer cells to regulate plasmin formation and cell migration. Mol. Oncol., 8: 483-507

  121. Li, S. and G.F. Hu, 2012. Emerging role of angiogenin in stress response and cell survival under adverse conditions. J. Cell. Physiol., 227: 2822-2826

  122. Lee, Y.H., C.W. Wei, J.J. Wang and C.T. Chiou, 2011. Rana catesbeiana ribonuclease inhibits Japanese Encephalitis Virus (JEV) replication and enhances apoptosis of JEV-infected BHK-21 cells. Antiviral Res., 89: 193-198

  123. Yadav, S.K. and J.K. Batra, 2015. Ribotoxin restrictocin manifests anti-HIV-1 activity through its specific ribonuclease activity. Int. J. Biol. Macromol., 76: 58-62

  124. Wang, H.X. and T.B. Ng, 2004. Purification of a novel ribonuclease from dried fruiting bodies of the edible wild mushroom Thelephora ganbajun. Biochem. Biophys. Res. Commun., 324: 855-859

  125. Zhu, M., L. Xu, X. Chen, Z. Ma, H. Wang and T.B. Ng, 2013. A novel ribonuclease with HIV-1 reverse transcriptase inhibitory activity from the edible mushroom Hygrophorus russula. Applied Biochem. Biotechnol., 170: 219-230

  126. Zhang, R., G. Tian, Y. Zhao, L. Zhao, H. Wang, Z. Gong and T.B. Ng, 2015. A novel ribonuclease with HIV-1 reverse transcriptase inhibitory activity purified from the fungus Ramaria formosa. J. Basic Microbiol., 55: 269-275

 
 
 
Insight Knowledge © 2018