17 Polynucleotide Phosphorylase

Polynucleotzde Phosphorylase U . 2. LITTAUER

H . SOREQ

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . A Occurrence and Intracellular Distribution . . . . . . . . . . . B . Purification . . . . . . . . . . . . . . . . . . . . . . . . . C . Molecular Weight of Whole Enzyme and Its Subunits . . . . . . D . Amino Acid Composition and Isoelectric Point . . . . . . . . . E . Immunological Analysis . . . . . . . . . . . . . . . . . . . F. Metal Ion Requirements . . . . . . . . . . . . . . . . . . . G . Stability and Sensitivity to Proteolytic Enzymes . . . . . . . . H . Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . I . Oligonucleotide Primers and Inhibitors . . . . . . . . . . . . J . Activators and Polyamines . . . . . . . . . . . . . . . . . . I11 . The Reactions Catalyzed . . . . . . . . . . . . . . . . . . . . A . Polymerization . . . . . . . . . . . . . . . . . . . . . . . B . Nucleoside Diphosphate-P, Exchange . . . . . . . . . . . . . C . Phosphorolysis . . . . . . . . . . . . . . . . . . . . . . . D . “Transnucleotidation” . . . . . . . . . . . . . . . . . . . . IV. Attributed Physiological Functions . . . . . . . . . . . . . . . . V. Research Applications . . . . . . . . . . . . . . . . . . . . . A . Polynucleotide Synthesis . . . . . . . . . . . . . . . . . . . B . Synthesis of Oligonucleotides with a Defined Sequence . . . . . C . Polymerization of Deoxyribonucleotides . . . . . . . . . . . . D . Conjugation to Insoluble Matrix . . . . . . . . . . . . . . . E . Synthesis of Radiolabeled Nucleotides and Fingerprinting of Oligonucleotides . . . . . . . . . . . . . . . . . . . . . . F. Synchronous Phosphorolysis as an Analytical Tool . . . . . . . G . Probe for the Regulatory Function of the 3’-OH Region of RNA . H . PNPase-Directed Labeling of the 3’-OH End of Polynucleotides .

.

518 519 519 520 522 523 524 525 525 528 529 529 530 531 534 535 537 537 539 539 543 545 546 547 548 550 553

517 THE ENZYMES. VOL . XV Copyright 0 1982 by Academic Press. Inc . All rights of reproduction in any form reserved ISBN C-IZ-l22715-4

518 1.

U. 2. LITTAUER AND H . SOREQ

Introduction

Polynucleotide phosphorylase (PNPase, polyribonucleotide :orthophosphate nucleotidyltransferase, EC 2.7.7.8) was discovered by Grunberg-Manago and Ochoa during the course of a study of the mechanism of biological phosphorylation in Azotobmter vinelundii (I -3). The enzyme catalyzes the reversible reaction formulated as follows [Eq. (l)]: Mg’+

~IPPN

(PN)n + nPi

(1)

Studies of the nature of ribonucleotide incorporation into nucleic acids led to a recognition of the same reaction inEscherichia coli extracts ( 4 , 5 ) .The enzyme was also isolated from Micrococcirs luteus (formerly classified as M. lysodeikticus) (6, 7), and subsequently has been shown to be widely distributed among bacteria (8). PNPase was the first enzyme to be discovered that can catalyze the formation of polyribonucleotides with a 3‘,5‘phosphodiester bond. In the forward reaction long polyribonucleotides are synthesized from various ribonucleoside diphosphates, with elimination of inorganic orthophosphate. Each of the four common ribonucleoside diphosphates can serve separately as a substrate for the polymerization reaction, leading to the formation of homopolymers. Polymerization of a mixture of nucleoside diphosphates that contain different bases results in the formation of a random copolymer, and the enzyme does not require a template and cannot copy one. Under suitable conditions the enzyme will also catalyze the elongation of a primer oligonucleotide with a free 3’-terminal hydroxyl group [Eq. (2)] as follows: R

+

n(ppN)

Mgz+

R(PN)~+ n P ,

(2)

where R represents the oligonucleotide primer, having at least two nucleoside residues and a free 3’-terminal hydroxyl group. In the reverse reaction, the enzyme catalyzes the breakdown of polyribonucleotides by phosphorolytic cleavage of the internucleotide I. 2. 3. 4. 5. 6. 7. 8.

Grunberg-Manago, M., and Ochoa, S. (1955). FP 14, 221. Grunberg-Manago, M., and Ochoa, S. (1955). JACS 77, 3165. Grunberg-Manago, M., Ortiz, P. J., and Ochoa, S. (1956). BBA 20, 269. Littauer, U. 2. (1956). FP 15, 302. Littauer, U. Z., and Kornberg, A. (1957). JBC 226, 1077. Beers, R. F., Jr. (1956). FP 15, 13. Beers, R. F., Jr. (1956). Nntirre (London) 177, 790. Grunberg-Manago, M. (1%3). Proyr. Nucleic Acid R r s . 1, 93.

5 19

17. POLYNUCLEOTIDE PHOSPHORYLASE

bonds. The phosphorolysis reaction proceeds in a stepwise fashion starting from the 3’-OH terminus of the polyribonucleotides to liberate NDPs. PNPase also catalyzes an exchange reaction between 32P-labeled inorganic phosphate and the P-phosphate of nucleoside diphosphates [Eq. (3)l Ribonu~leoside-”P-~~P + .’‘P

Mgs ‘

R i b o n u c l e ~ s i d e - ~ ~ P+- ~31P ~P

(3)

All these reactions have served as a basis for the assay of enzyme activity. Several excellent review articles summarize the extensive work carried out with this enzyme (8-15). In the following sections we summarize the current knowledge regarding the enzyme, its purification, properties, and the various reactions it catalyzes. In particular, we will emphasize the wide range of research applications that are in use with this enzyme. II.

A.

Properties

OCCURRENCE

AND

INTRACELLULAR DISTRIBUTION

PNPase is widely distributed among different aerobic, anaerobic, and halophilic bacteria [cf. Ref. ( I S ) ] . It was also isolated from Brevibacterium (161, B. stearc~tliermophilirs(17), Thermris crqirnticus (or Tliermus thermophilw) (17. 18), and the photosynthetic bacterium Rhodospirillum rubrum ( 1 9 ) . Achromobacter sp. KR 170-4 (20) and the bacteroid form of Rhiwbium meliloti (21) seem to be relatively rich sources for the enzyme. The properties of PNPase seem to differ somewhat in various bacterial species. 9. Steiner, R. F., and Beers, R. F., Jr. (1961). “Polynucleotides, Natural and Synthetic Nucleic Acids.” Elsevier, Amsterdam. 10. Grunberg-Manago, M. (1961). “The Enzymes,” 2nd ed., Vol. V, p. 257. 11. Grunberg-Manago, M. (1962). ARB 31, 301. 12. Grunberg-Manago, M. (1963). Progr. Biopfivs. M o l w . B i d . 13, 175. 13. Singer, M. F. (1966). I n “Procedures in Nucleic Acid Research” (G. L. Cantoni and D. R. Davies, eds.), p. 245. Harper and Row, New York. 14. Thang, M. N. (1969). Bull. Soc. Cliim. B i d . 51, 1407. 15. Godefroy-Colburn, T., and Grunberg-Manago, M. (1972). “The Enzymes,” 3rd ed., Vol. 7, p. 533. 16. Yang, H. H.. Thayer, D. W., and Yang, S. P. (1979). Appl. Environ. Microhiol. 38,

143. 17. Wood, J. N., and Hutchinson, D. W. (1976). N/rc./ric Acids R e s . 3, 219. 18. Hishinuma, F., Hirai, K., and Sakaguchi, K. (1977). EJB 77, 575. 19. Soe, G., and Yamashita, J . (1980). J B 87, 101. 20. Rokugawa, K., Katoh, Y., Kuninaka, A., and Yoshino, H. (1975). A g r . B i d . Chrm. 39, 1455. . 21. Hunt, R. E., and Cowles, J. R. (1977). C O NMicrobid. 102, 403.

520

U. Z. LITTAUER AND H. SOREQ

PNPase is found in the soluble fraction of many bacterial cells ( 2 2 , 2 3 ) . Ribosomes ofE. coli contain some enzyme activity; however, most of the PNPase can be removed by repeated washing. About 10% of the total activity remains attached to washed ribosomes, probably bound to mRNA ( 2 2 ) . Some activity is also found in membrane vesicles isolated from E. coli cells ( 2 4 ) . In Streptococcus fuecalis ( 2 5 ) , S . pyogenes ( 2 6 ) , and Halohacterium cutirubrirm ( 2 7 ) , however, the enzyme is found in the cell membranes. PNPase has also been detected in wheat roots (28), and partially purified from healthy and tobacco mosaic virus (TMV)-infected tobacco leaves. However, its localization within the plant cell is uncertain (29). Partial purification of PNPase from the blue-green alga Anacysris nidufans has also been described (30). Similar activities have been reported in animal cells [cf. Refs. (15, 31)], although the results could be due to a combination of other enzymes ( 3 2 ) . Enzymatic activity that catalyzes the phosphorolysis of polyribonucleotides to NDPs has been partially purified from guinea pig liver nuclei. Unlike bacterial PNPase, the animal enzyme does not appear to catalyze the synthesis of polynucleotides (33). In addition to being associated with the nuclear membrane from rat liver cells (34, 35), PNPase activity is associated with the inner membrane of their mitochondria ( 3 4 ) . Enzymatic activity has also been detected in the endoplasmic reticulum of ribosome fraction from regenerating liver cells (35-37). B. PURIFICATION Bacterial PNPase has been purified from a wide variety of sources [reviewed in Ref. (15)l. Improved isolation procedures have increased 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37.

Kimhi, Y., and Littauer, U. Z. (1967). Biochemistry 6, 2066. Owen, P., and Salton, M. R. J. (1977). J B 132, 974. Owen, P., and Kaback, H. R. (1979). Biochemistry 18, 1413. Abrams, A., and McNamara, P. (1962). JBC 237, 170. Kessler, R. E . , and van de Rijn, I. (1979).Infect. Zmmrrn. 26, 892. Peterkin, P. I., and Fitt, P. S. (1971). BJ 121, 613. Kessler, B., and Chen, D. (1%4). BBA 80, 533. Brishammar, S. , and Juntti, N. (1974). ABB 164, 224. Capesius, I., and Richter, G . (1967). Z.Nfiturfvrschg. 22b, 204. Fitt, P. S . , and See, Y.P. (1970). BJ 116, 309. Smellie, R. M. S. (1963). Progr. Nucleic Acid Res. 1, 27. See, Y.P., and Fitt, P. S. (1970). BJ 119, 517. See, Y.P., and Fitt, P. S. (1971). FEBS Lett. 15, 65. Delvig, A. A. (1978). Biokhimiu 43, 579. Delvig, A. A., and Mardachev, S. R. (1975). Biokhimiu 40, 1246. Delvig, A. A., Tarasov. A. P.. and Debov, S. S. (1976). Biokhimiu 41. 2201.

17. POLYNUCLEOTIDE PHOSPHORYLASE

52 1

both the yield and purity of the enzyme. Essentially homogeneous preparations have been obtained from E. coli (38-42) M . luteus (43, 44), A . vineiondii (45, 461, C . pedringens (47), B . srrarothermoplzilus ( 1 7 ) , Thermus thermophilus (17, 18), and Rhodospirillum rubrum (19). Afinity chromatography on columns of poly(A)-Sepharose (48),p-aminophenyl oligo(dT)-Sepharose (491, RNA-Sepharose (42), poly(1)-agarose (50), Blue-Dextran-Sepharose (50, 51 ) and poly(U)-Sepharose (44) have yielded substantial purification of the enzyme. The effectiveness of these methods depends on prior removal of nucleic acid contaminations from the crude enzyme preparations. Phenylmethylsulfonyl fluoride has been included in solutions used for PNPase purification because of the sensitivity of the enzyme to proteolytic degradation (41, 44). Purified enzyme preparations from M. luteus (44) and B. stenrothermophilus ( 1 7 ) are virtually free of contaminating nucleic acids. Escherichia coli PNPase purified by different procedures contains low levels of bound oligonucleotides (39, 41, 42), as does the enzyme from R. rubrum (19). Most of the bacterial PNPase preparations are primer-independent forms and catalyze de novo polymerization in a processive fashion. With these enzyme forms the rate of the polymerization reaction is only slightly stimulated by oligonucleotides. Early purification of M. lufeus PNPase yielded primer-dependent preparations, in which the polymerization reaction is almost completely dependent on the presence of oligonucleotides ( 1 3 , 5 2 ) . However, with subsequent batches of cells, only the primer-independent form (form I) could be obtained (44, 5 3 ) . Primer-dependent form (form T) can be derived from the inde38. Williams, F. R., and Grunberg-Manago, M. (1964). BBA 89, 66. 39. Kimhi, Y.,and Littauer, U. Z. (1968). JBC 243, 231. 40. Kimhi, Y.,and Littauer, U. Z. (1968). “Methodsin Enzymology,” Vol. XIIB, p. 513. 41. Portier, C., van Rapenbusch, R., Thang, M. N., and Grunberg-Manago, M. (1973). EJB 40, 77. 42. Soreq, H., and Littauer, U. Z. (1977). JBC 252, 6885. 43. Letendre, C. H., and Singer, M. F. (1975). Nucleic Acids Res. 2, 149. 44. Barbehenn, E. K., Craine, J. E., Chrambach, A., and Klee, C. B. (1982). JBC 257, 1007.

45. Gajda, A. T., Zaror de Behrens, G., and Fitt, P. S. (1970). BJ 120, 753. 46. Mii, S. (1977). J B 81, 899. 47. Guissani, A. (1978). Biochimie 60,755. 48. Lehrach, H., and Scheit, K. H. (1972). Hoppe Seyyler’s Z. Physiol. Chem. 353, 731. 49. Smith, J. C., and Eaton, M. A. W. (1974). Niicleic Acids Res. 1, 1763. 50. Drocourt, J. L., Thang, D. C., and Thang, M. N. (1978). EJB 82, 35.5. 51. Thang, M. N., Drocourt, J. L., Chelbi-AEx, M. K., Thang, D. C., Lubochinski, J., Ruet, A., Sentenac, A., Gangloff, J., and Dirheimer, G. (1979). Cmlloq. Inserm. Afiniry Chromatogr. 86, 303. 52. Singer, M. F., and Guss, J. K. (1962). JBC 237, 182. 53. Klee, C. B. (1967). JBC 242, 3579.

522

U . Z. LITTAUER AND H. SOREQ

pendent form by limited tryptic digestion (54,55). Following trypsin digestion, A. vinelandii PNPase also develops primer requirement (45). PNPase preparation purified from B. stearothermophilus (I 7) and T. thermophilus (18) show primer dependency probably due to endogenous proteolysis.

c.

MOLECULAR WEIGHT OF WHOLE ENZYME ITS SUBUNITS

AND

The molecular weight of the whole enzyme has been determined by sedimentation equilibrium, gel filtration, sucrose gradient centrifugation, and gel electrophoresis under nondenaturing conditions. In the latter method, enzyme activity can be visualized after electrophoresis by incubating the gels in the presence of ADP and Mg‘+, followed by staining the poly(A) formed in situ with acridine orange in the presence of lanthanum chloride (56). Other in sitir methods for visualizing active enzyme molecules have also been published (23, 45, 57). The physicochemical properties of E. coli form A (4f142,58) and M. luteus form I PNPase are similar (44).The molecular weight of purified E. coli PNPase determined by sedimentation equilibrium ranges between 230,000 +- 20,000 (42) and 216,000 I+_ 20,000 (4f), as compared to 237,000 24,000 for the M. iuteus enzyme (44). A value of 252,000 has been calculated from a Stokes radius of 6.4 nm and a sedimentation constant of 8.9 S for theE. coli enzyme (58). The observed frictional ratio is 1.52 (58). The E. coli enzyme is composed of three identical subunits of a molecular weight ranging between 84,000-95,000 ( 4 I , 42, 48, 58, 5 9 ) . Support for the a3 structure of E. coli PNPase arises from ultrastructural observations. Under the electron microscope, the enzyme appears as a triangle with a central hole. The diameter of these molecules was calculated to be 85 A (60). In crude cell extracts PNPase displays microheterogeneity . Sucrose gradient sedimentation, gel filtration, and gel electrophoresis all show the presence of higher level components ( M , 39, 4f.43, 44,54,58). These forms arise from the association of an additional polypeptide subunit (41. 58) or the presence of bound nucleic acids that

*

54. Klee, C. B. (1969). JBC 244, 2558. 55. Klee, C. B. (1971). I n “Procedures in Nucleic Acid Research” ( G . L. Cantoni and D. R. Davies, eds.), Vol. 2, p. 896. Harper and Row, New York. 56. Thang, M. N . , Thang, D. C., and Leautey, J. (1967). C . R . Acad. Soc. ( P a r i s ) 265, 1823. 57. Fitt, P. S ., Fitt, E. A., and Wille, H. (1968)’. BJ 110, 475. 58. Portier, C. (1975). EJE 55, 573. 59. Portier, C. (1975). FEBS Lett. 50, 79. 60. Valentine, R. C., Thang, M. N . , and Grunberg-Manago, M. (1969). J M B 39, 389.

17.

POLYNUCLEOTIDE PHOSPHORYLASE

523

induce a conformational change in the enzyme (44). PNPase from E. coli can be isolated in two active forms, A or B, having molecular weights of 252,000 and 365,000, respectively. The A form has an a3type structure, whereas the B form has two types of chains, a (MW 86,000) and /3 (MW 48,000). The exact proportion of the a and p subunits is not yet clear, and PNPase B form has been assigned a structure of a& (or a3Pn). The B form is obtained by keeping the ionic strength at 0.2 M during the purification of the enzyme on a Sephadex G-200 column, whereas at lower salt concentrations the /3 subunit tends to dissociate and the enzyme reverts to the A form. All the catalytic activity of PNPase resides in the a subunits, whereas the p subunit is inactive and does not alter the enzymatic properties of the whole enzyme. The role of the p subunit therefore remains to be determined (58). In addition to the main 252,000 MW form, E . coli B and K12 extracts contain 25 and 596, respectively, of a low molecular weight (100,000) PNPase. The 100,000 MW form catalyzes the phosphorolysis reaction but is unable to catalyze the polymerization of NDPs. The 100,000 form differs from the main 252,000 MW enzyme in that it can only phosphorolyze short-chain polymers and requires higher Mg2+ ion concentrations. PNPase preparations from E. coli Q13 and 1 1 13 mutants are particularly rich in this defective enzyme, and about 80% have a molecular weight of 100,000. In addition, about 20% of the mutant PNPases have a molecular weight of about 200,000. Unlike the 100,000 MW form, or the wild-type enzyme, this additional form requires Mn2+for NDP polymerization and has a higher K , for poly(A) phosphorolysis (61). Clostridirrm per-ingens PNPase also appears in two forms, a& and a3, with molecular weights similar to that of the E. coli enzyme (47). PNPase from A . vinelmdii has an apparent molecular weight of 200,000 (46). But the R. ruhrum enzyme has an MW of 160,000, and appears as a dimer of two subunits of 76,000 (19). PNPase from B . stearothermophilus (17)is a tetramer of 51,000, and that from T. thermophilus (18) shows three subunits of 92,000, 73,000, and 35,000, which may result from limited proteolysis of the enzyme.

D. A M I N OACID COMPOSITION

AND

ISOELECTRIC POINT

The amino acid composition of the E. coli PNPase (42, 62) is similar to that of the M . lrrteirs enzyme (43.44). Although there are some differences in the reported cysteine and tryptophan values for the E. coli enzyme (42, 61), it is likely that it contains 3 Cys and 3 Trp residues per mole of subunit 61. Thang, M. N . , Thang, D. C . , and Grunberg-Manago, M. (1969) EJB 8, 577. 62. Portier, C. (1975). Biocliirnie 57, 545.

524

U. Z. LITTAUER AND H. SOREQ

of 84,000 MW, as does the M. lureus enzyme (44). Of the 3 Cys residues of the E. coli subunit, only one group is exposed, and is found to react with dithionitrobenzene, whereas 2 groups are “masked” and react only after denaturation with 1% sodium dodecyl sulfate (42). This property may explain the insensitivity of the enzyme to -SH reagents (39). The UV difference spectrum of the M. luteus enzyme suggests that 6 Tyr residues (out of 17), and perhaps 1 Trp residue (out of 3), are buried in the interior of the protein and become exposed upon treatment with 6 M guanidineHCl(44). One Trp and three Tyr residues are lost during the conversion of M. luteus form I to form T, which may explain why form T has lost both the ability to bind oligonucleotides with high affinity and to catalyze de novo synthesis of poly(A) (44). The a and /3 subunits of E. coli PNPase appear to be unrelated to each other, and differ in their amino acid composition as well as in their cyanogen bromide cleavage peptides (62). The N-terminal amino acid sequence of the E. coli a form suggests that the a chains are all identical and terminate with Met-Leu-?-Pro-Phe (62). Methionine is also the only amino acid found at the N-terminus of B. srenrothermophilus enzyme (17). In contrast, neither the I nor the T form of the M. luteus enzyme contains detectable free amino end groups. Since the primer-dependent T form is obtained by limited trypsin digestion, it is suggested that proteolysis removes a peptide at the carboxy end of the molecule (44). In situ staining of isoelectric focusing gels revealed an isoelectric point of 6.1 for PNPase from E. coli (42) and M . luteus (44), whereas the enzyme preparations isolated from B. stearothermophilus (17) and T. thermus (18) focus at pH 4.1 and 4.3, respectively.

E.

IMMUNOLOGICAL ANALYSIS

Antibodies against purified PNPase from E. coli B were shown to react with the enzyme in a double diffusion test and in immunoelectrophoretic analysis. The enzyme, complexed with its antibodies, retains its polymerization properties, and the antigen-antibody complex can be visualized by autoradiography of the polynucleotide formed in situ by the enzyme (63). Double precipitation bands were obtained with enzyme purified according to Williams and Grunberg-Manago (38, 63). However, rabbit antibodies elicited against homogeneous E. coli PNPase, following the affinity chromatography step, displayed a single precipitation band (42). No serological relationship exists between E. coli PNPase and either the core enzyme or the L+ subunit of E. coli RNA polymerase (42), in 63. Uriel. J., Thang, M. N., and Berges, J. (1969). FEES Lett. 2, 321.

525

17. POLYNUCLEOTIDE PHOSPHORYLASE

contradistinction to an earlier suggestion (64). The ribosomal S 1 protein, which contaminates E. coli PNPase, is also unrelated to PNPase (42). F. METALION

REQUIREMENTS

Many studies indicate that Mg" is required for the reactions catalyzed by PNPase and that it can be partially replaced by Mn'+ (5, 15).Free Mgz+ M (65). Other cations, ions bind to E. coli PNPase with a K,,, of 5 x such as Co2+,Ni", Cd2+,Cu2+,and Zn2+,but not Ca'+, may also replace Mg2+in PNPase reactions, although with quite different efficiencies (19, 39, 66). Polymerization of GDP with E. coli PNPase, however, proceeds efficiently in the presence of Mn" at 60" (67). The polymerization reaction with a mutant PNPase from E. coli 413 requires Mn2+rather than Mg'+ (68), and Mn2+will stimulate more efficiently than Mg'+ the polymerization reaction with PNPase from Achromohucter (20). If, indeed, PNPase plays a role in the nucleolytic degradation of RNA (69), the inability of Ca2+to replace Mg2+in the phosphorolysis reaction with E. coli PNPase may partially contribute to the protective effect that Ca2+exerts in vitro on various types of mRNA (70). However, at low Ca2+concentration, of about 5 p M , there is a threefold activation of the polymerization reaction with B. steurothermophilus enzyme (17). At suboptimal Mg" concentrations, both the formation of polymers from NDPs and the NDP-Pi exchange reaction occur only after an initial lag period. In the presence of polynucleotides or short oligonucleotides, this lag period is almost abolished (38, 71-73).

G.

STABILITY A N D SENSITIVITY TO PROTEOLYTIC

ENZYMES

Purified E. coli PNPase is unstable above 55", and is rapidly and irreversibly inactivated at 65" (15, 39, 42). The M. I u t e ~ s(74) and the C. per64. Ohasa, S . , Tsugita, A., and Mii, S . (1972). Nature N e w B i d . 240, 39. 65. Williams, F. R . , Godefroy, T., Mery, E . , Yon, J . , and Grunberg-Manago, M. (1964). BBA 80, 349. 66. Babinet, C., Roller, A., Dubert, J. M., Thang, M. N., and Grunberg-Manago, M. (1965). BBRC 19, 95. 67. Thang, M. N., Graffe, M . , and Grunberg-Manago, M. (1965). EEA 108, 125. 68. Hsieh, W. T., and Buchanan, J. M. (1967) PNAS 58, 2468. 69. Kaplan, R . , and Apirion, D. (1974).JBC 249, 149. 70. Cremer, K . , and Schlessinger, D. (1974). JEC 249, 4730. 71. Ochoa, S . , and Mii, S. (1961). JBC 236, 3303. 72. Mii, S . , and Ochoa, S. (1957). EBA 26, 445. 73. Singer, M. F., Heppel, L. A., and Hilmoe, R. J. (1957). BBA 26, 447. 74. Brenneman, F. N., and Singer, M. F. (1964). EBRC 17, 401.

526

U . Z. LITTAUER AND H. SOREQ

fringens enzyme (75) are less stable than E. coli PNPase. The enzyme is stabilized against heat inactivation by the presence of NDPs, but not by NMPs, NTPs, or DNA. Substrate oligonucleotides with free 3'-OH termini can also exert this protective effect, whereas oligonucleotides with blocked 3' ends do not affect the rate of heat inactivation (76). Heatdenatured E. coli PNPase can be renatured. Following heating at 100" for 1 min the precipitate is dissolved in 6 M guanidine-HC1followed by dialysis. About 25-30% of the original enzyme activity is recovered by this procedure, and the reassociated enzyme reverts to its original quaternary as structure (41). High concentrations (>3.0 M) of urea have also been shown to cause inactivation of E. coli PNPase. In this case as well, the presence of substrates protects the enzyme against the inactivation process (77). PNPase is sensitive to proteolytic digestion. Earlier studies revealed differences in subunit structure and catalytic properties of the enzyme when isolated from various bacterial sources. It now appears that these differences mainly result from endogenous proteolytic digestion in various enzyme preparations, and that the properties of the various intact enzyme preparations are similar. Degradation by endogenous proteases of PNPase from M. luteus (53),A . ngilis (45), C. pegringens (47, 78), and E. coli (79, 80), or degradation with chymotrypsin or trypsin (45, 47, 54, 57, 79-81) yield very close gel electrophoretic patterns. Limited proteolysis of the enzyme supports the view that the catalytic center and the polynucleotide binding subsite (82-86) are distinct and dispersed over the enzyme surface. Storage of E. coli PNPase for extended periods at 4" results in limited proteolysis of the enzyme. The proteolyzed PNPase has a reduced molecular weight of 175,000 with an a$ structure (a' = 65,000). The endogenous proteolysis induces changes in both the phosphorolysis and polymerization reactions (79, 80). The K , 75. Fitt, P. S . , Dietz, F. W., Jr., and Grunberg-Manago, M. (1968). BBA 151, 99. 76. Lucas, J . M., and Grunberg-Manago, M. (1964). BBRC 17, 395. 77. Harvey, R. A . , Godefroy, T., Lucas-Lenard, J., and Grunberg-Manago, M. (1967). EJB 1, 327. 78. Guissani, A., and Grunberg-Manago, M. (1969). BBRC 35, 131. 79. Thang, M. N . , Dondon, L., and Godefroy-Colburn, Th. (1971). Biochimie 53, 291. 80. Guissani, A . , and Portier, C. (1976). Nucleic Acids Res. 3, 3015. 81. Fitt, P. S . , and Wille, H. (1969). BJ 112, 497. 82. Chou, J . Y . , and Singer, M. F. (1970). JBC 245, 995. 83. Thang, M. N . , Guschlbauer, W., Zachau, H. G . , and Grunberg-Manago, M. (1967). J M B 26, 403. 84. Thang, M. N . , Harvey, R. A., and Grunberg-Manago, M. (1970). JMB 53, 261. 85. Godefroy, T. (1970).EJB 14, 222. 86. Chou, J. Y . , Singer, M. F., and McPhie, P. (1975). JBC 250, 508.

17. POLYNUCLEOTIDE PHOSPHORYLASE

527

to for a polynucleotide in the phosphorolysis reaction shifts from M , indicating that proteolysis causes a loss of the polynucleotide binding site. The proteolyzed enzyme shows a much more stringent requirement for an oligonucleotide primer in the polymerization reaction but is not stimulated by polynucleotides. Because of the loss of polynucleotide binding sites, phosphorolysis of poly( A),U with the proteolyzed enzyme proceeds with a partially nonprocessive mechanism, as opposed to the processive phosphorolysis displayed by native enzyme. It has been assumed (80) that the polymerization mechanism as well will no longer be purely processive, and that the mean length of the polymers synthesized will be shorter than that observed for polymers obtained with native enzyme. The proteolyzed enzyme also fails to bind to polynucleotide-agarose or Blue Dextran-agarose columns (SO, 5 1 ) . As with native enzyme (87), phosphorylation of proteolyzed E. coli PNPase by cyclic AMP-dependent protein kinase can replace the stimulating effect of oligonucleotides in the polymerization reaction proteolyzed (79). Similar changes in the properties of E . coli PNPase were also produced by incubating the enzyme with isolated bacterial proteases (79, 88). Native PNPase from M . luterrs is primer-independent, catalyzes de n o w polymerization in a processive fashion, and is only slightly stimulated by oligonucleotides. Limited trypsin digestion of the native enzyme alters its polymerization activity without affecting its ability to phosphorolyze polynucleotides. The trypsinized enzyme (form T) catalyzes the elongation of primer by a random mechanism and is stimulated up to 20-fold by oligonucleotides (54, 57, 89 -91 ). Restoration of primerindependence to form T can be obtained by treatment with Pmercaptoethanol. Reconversion to primer dependence is achieved by reaction with sulfhydryl inhibitors, suggesting that the alteration in the enzyme properties is correlated with the modification of sulfur-containing amino acids (54, 92). Enhancement in primer requirement also appears in PNPase from Azorobacter tinehzdii upon mild treatment with trypsin or aging of the enzyme (45, 49, 93). Restoration of the reduced activity, but not the loss of primer requirements, is caused in this case as well by P-mercaptoethanol. 87. Thang, M. N . , and Meyer, F. (1971). FEES Lc,tr. 13, 345. 88. Regnier, Ph., and Thang, M. N. (1972). Biocliirnie 54, 1227. 89. Moses, R . E., and Singer, M . F. (1970). JBC 245, 2414. 90. Klee, C. B . , and Singer, M. F. (1968). JBC 243, 923. 91. Fitt, P. S . , and Fitt, E. A. (1967). BJ 105, 25. 92. Klee, C . B . , and Singer, M. F. (1968). JBC 243, 5094. 93. Gajda, A. T., and Fitt, P. S. (1969). BJ 112, 381.

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U . Z. LITTAUER AND H . SOREQ

PNPase from C. perfringens is highly susceptible to proteolysis and is obtained as a mixture of variable proportions of native and proteolyzed forms. Under the action of either endogenous proteases or trypsin, two enzymatic forms are obtained that differ in their catalytic properties from each other and from the initial enzyme. One of the proteolyzed species catalyzes polymerization only in the presence of poly(A) or polylysine, whereas the other phosphorolyzes oligonucleotides but not polynucleotides (47). In contrast to the native enzyme, the proteolyzed enzyme requires P-mercaptoethanol and polylysine for efficient polymerization activity (47, 7 8 ) .

H. INHIBITORS Several chemical agents have been shown to block the catalytic activity of PNPase from various biological sources. Some of these, such as 6-azauridine or 5-fluorouridine diphosphates (94), as well as phosphonic acid analogs of ADP (95, 96) or analogs produced by periodate oxidation (97),appear to react with the active site and inhibit the exchange, the phosphorolysis, and the polymerization activities of the enzyme. Inhibitory reaction has also been noticed for deoxynucleoside diphosphates (98, 99). Other inhibitors, such as acridine orange, appear to inhibit the polymerization reaction via their interaction with the primer oligonucleotide (100, f 01). The catalytic activity of PNPase from B. amyloliquefuciens has been reported to be inhibited by heparin, rifamycin SV, and synthetic polynucleotides (102); the polymerization reaction catalyzed by E. coli PNPase is effectively inhibited by oligophosphates of pyridoxal, the percentage of inhibition being higher with longer chains of phosphate moieties bound to the pyridoxal core (103). 94. Skoda, J., Kara, J . , Sormova, Z., and Sorm, F. (1959). BBA 33, 579. 95. Simon, L. N . , and Myers, T. C. (l%l). BBA 51, 178. %. Godefroy-Colburn, T., and Setondji, J. (1972). BBA 272, 417. 97. Smrt, J., Mikhailov, S. N . , Hynie, S. , and Florentev, V. L. (1975). Collect. Czech. Chem. Commun. 40, 3399. 98. Lucas-Lenard, J . , and Cohen, S. S. (1966). BBA 123, 471. 99. Bon, S., Godefroy, T., and Grunberg-Manago, M. (1970). EJB 16, 363. 100. Beers, R. F.,Jr.. Hendley, D. D., and Steiner, R. F. (1958). Nature (London) 182, 242. 101. Beers, R. F., Jr. (1%0). JBC 235, 726. 102. Erickson, R. J . , and Grosch, J. C. (1977). J B 130, 869. 103. Mamaeva, 0. K., Karpeiskii, M. Ya., Karpeiskii, A. M., and Bibilashvili, R. Sh. (1979). Molek. B i d . 13, 811.

17. POLYNUCLEOTIDE PHOSPHORYLASE

I.

529

OLIGONUCLEOTIDE PRIMERS A N D INHIBITORS

The de n o w polymerization of NDPs, particularly at low Mg2+concentration, is preceded by a lag period, which may be overcome by the addition of polynucleotides or short oligonucleotide primer molecules with a free 3’-hydroxyl group (38,39, 71-73). These primers also accelerate the exchange of phosphate moieties by the enzyme (39, 73, 104). The oligonucleotide primers have been shown to be incorporated into the polymer synthesized by PNPase from M. luteus (105), and their effect was found to be maximal in the polymerization of GDP, which proceeds with difficulty and at a slow rate in the absence of such primers (106). When blocked with a 3’-terminal phosphate moiety, oligonucleotides act as inhibitors of the polymerization and the exchange reactions (71, 72, 107). The inhibition is temperature-dependent and may be overcome by addition of a complementary polynucleotide, which hybridizes with the oligonucleotide inhibitor and prevents its binding to the enzyme (108). The strong binding of blocked polynucleotides has been exploited to develop affinity chromatography procedures to purify the enzyme (42 ).

J. ACTIVATORS A N D POLYAMINES The polymerization reaction catalyzed by PNPase has been reported to be activated by several agents. Potassium, sodium, and lithium salts have been shown to affect the K, values of the M. luteus enzyme (109). A basic polypeptide that enhances the ADP-Pi exchange reaction has been isolated from E. coli extracts (5, 39). In the presence of this heat-stable activator, the optimal Pi concentration for ADP-Pi exchange shifts from 2 to about 0.65 mM. At low phosphate concentrations, the activator causes up to 3- to 6-fold stimulation of the exchange reaction withE. coli PNPase, but has no effect on the rate of polymer formation or the phosphorolysis of poly(A). The activation of the exchange reactions with NDPs other than ADP is much lower than with ADP. Spermine and spermidine (0.1- 1.0 mM) also activate the ADP-Pi exchange (twofold), whereas poly-L-lysine and poly-L-ornithine M ) hardly affect the reaction, and at higher concentrations cause inhibition (89). A basic protein from A. vinelandii 104. 105. 106. 107. 108. 109.

Beers, R . F., Jr. (1961). JBC 236, 2703. Singer, M . F., Heppel, L. A . , and Hilmoe, R . J. (1960). JBC 235, 738. Brenneman, F. N . , and Singer, M. F. (1964). JBC 239, 893. Beers, R . F., Jr. (1959). Nufitre (London) 183, 1335. Heppel, L . A . (1963). JBC 238, 357. Beers, R . F., Jr. (1957). Nutitre (London) 180, 246.

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U. Z. LITTAUER AND H. SOREQ

causes a lag phase in the NDP polymerization reaction. Preferential repression of polymerization of UDP is observed with polylysine and of ADP with polyarginine. A lag phase is also caused by polylysine in the ADP-Pi exchange reaction with the A . vinekcindiii enzyme (46). The stimulating activity of polylysine and other polyamines on the ADP polymerization with proteolyzed C. perfringens enzyme has been noted (47, 75, S l ) , and is probably due to charge effects on the purified protein (1 10). A different mode of activation of the polymerization reaction is exerted by acridine orange, which forms complexes with the newly synthesized polynucleotides in the reaction mixture and, changing the equilibrium constants, drives the reaction toward further polymerization (100. 101). Yet another mechanism of activation has been observed with ATP, which improves the yields of polynucleotides synthesized with crude extracts from Azotobcicter vinelcindii (1 11). Activation by ATP has been suggested to function via the phosphorylation of PNPase by CAMP-dependent protein kinase (87). It should, however, be noted that these studies were mostly carried out with partially purified enzyme preparations, and that the effects observed could result from combined changes in the activities of contaminating enzymes. 111.

The Reactions Catalyzed

PNPase catalyzes the phosphorolysis of long-chain polynucleotides in a processive mechanism [also denoted progressive, see Ref. (IS)],whereby the enzyme does not dissociate from the polymeric substrate during the degradation process. Thus, the enzyme appears to degrade one polymer chain to completion prior to releasing a small resistant oligonucleotide and initiating phosphorolysis of another chain (15, 83, 90). In contrast, short oligonucleotides are degraded by a random nonprocessive mechanism [also denoted synchronous, see Ref. ( I S ) ] in which the enzyme dissociates from the substrate after hydrolysis of each nucleotide (82, 112). In addition, PNPase catalyzes the de nuvo polymerization of NDPs to polynucleotides in a processive mechanism, whereas elongation of oligonucleotide primers may occur by a nonprocessive mechanism [cf. Ref. (15)]. The diversity of mechanisms seems to be due to the existence of two classes of 110. Fitt, P. S., and Wille, H. (1969). BJ 112, 489. 1 1 1 . Shiobara, Y., and Itagaki, K. (1963). J B 54, 317. 112. Singer, M. F., Hilmoe, R. J., and Grunberg-Manago, M. (1960). JBC 235, 2705.

17. POLYNUCLEOTIDE PHOSPHORYLASE

53 I

binding sites in the molecule (84-86, 113-115). The first site, subsite I, is the catalytic center of the molecule and includes the mononucleotide, inorganic phosphate, and oligonucleotide binding domains. Subsite I binds the 3’-OH terminus of the growing polynucleotide or oligonucleotide. When polynucleotides are long enough, they can reach a second site, subsite 11, which is not involved in the phosphorolysis of oligonucleotides. Subsite I1 probably includes several polynucleotide binding domains (116). It has also been suggested that subsite I1 includes a lysinerich area that may act as a regulatory site (46). The residence time of the polymer in subsite I is short, whereas in subsite I1 it is long and corresponds to a very strung affinity of the enzyme for polynucleotides (lO-*M to M ) . This dual attachment to the enzyme allows long polynucleotides to snap back to a reactive position after removal of one nucleotide residue and thereby be degraded in a processive manner. Thus, binding to subsite I1 is responsible for the marked enhancement in the binding of polynucleotides. Oligonucleotides, not being anchored at subsite 11, are lost into the solution after the reaction and released from subsite I, and may then be replaced by another substrate molecule, leading to a nonprocessive random mechanism (15, 85, 86, 116). Limited proteolytic degradation (80, 1/.5), as well as linking the E. coli enzyme to BrCNactivated Sepharose ( I 17), affects mainly the polynucleotide binding domains of subsite 11, and results in the loss of complete processiveness and the decrease of affinity for polynucleotides. The active center is preserved, however, indicating that subsite I is in some way hidden. These findings are in agreement with the a3 subunit structure, which envisages the enzyme to have a triangle profile with a central hole in which the active center might be located (59, 60, 117). A.

POLY M E RIZ AT I ON

1. Initiritioti of de Novo Synthesis

PNPase catalyzes de novo synthesis of polynucleotides. The mechanism of formation of the first internucleotide bond is still unclear and probably involves the reaction between two NDP molecules, out of which one serves as the accepting 5’ terminus. Analysis of the newly synthe113. 114. 115. 116. 117.

Kaufmann, G., and Littauer, U. 2. (1969). FEES L d r . 4, 79. Chou, J . Y., and Singer, M. F. (1970). JBC 245, 1005. Guissani, A. (1977). EJE 79, 233. Godefroy, Th., Cohn, M., and Grunberg-Manago, M. (1970). EJB 12, 236. Vang, N . H . , Drocourt, J. L., and Thang, M . N. (1979). BBRC 90, 606.

532

U. Z. LITTAUER AND H. SOREQ

sized polynucleotides reveals that they contain, at the 5’ terminus, a monophosphate group rather than the expected pyrophosphate group (1 18). It is possible that a 5’-pyrophosphate terminus is initially formed, followed by removal of the @phosphate (or NDP) from the 5’ terminus at a later stage in the reaction. A novel mechanism suggests the transfer of the p-phosphate of ADP such that the AMP product formed can be positioned on PNPase as the 5’-monophosphate terminus of the nascent poly(A) chain. This transfer could depend on the deoxyadenylate kinase activity that is associated with PNPase from M. luteus (1 19). It should be noted that ApA and pApA do not undergo phosphorolysis, and accumulate as resistant end products of poly(A) phosphorolysis (1 14, 120-122). This would imply that the initiation of de nova polymerization involves an initial irreversible step. 2 . Elongation The processive elongation of polynucleotides by PNPase proceeds at a linear rate and then reaches a plateau. The polymers formed are of high molecular weight and are homogeneous in size; no intermediate oligonucleotides are formed (84). On the basis of kinetic analysis, the presence of a transient intermediate polynucleotide of high molecular weight, which is subsequently degraded to an equilibrium mixture of short oligonucleotides, has been proposed for primer-dependent PNPase (123). The complex enzyme-polynucleotide does not dissociate during the elongation process, even when unfavorable substrates such as GDP are polymerized at high temperature and in the presence of MnZ+ (67). PNPase utilizes the Sp diastereomer (exoisomer) of NDPS as a substrate, whereas the Rp isomer is a competitive inhibitor. During polymerization an inversion occurs in the configuration of the phosphorous bond into the Rp type (endoisomer), as was shown by high performance liquid chromatography of uridine 2’ ,3’-cyclic phosphorothioate, enzymatically obtained from copolymers of UDP with adenosine 5’-0-(1thiodiphosphate) (124). 118. 119. 120. 121. 122. 825. 123. 124.

Harvey, R. A., and Grunberg-Manago, M. (1966). BBRC 23, 448. Craine, J. E., and Klee, C. B. (1976). Nucleic Acids Res. 3, 2923. Singer, M. F. (1958). JBC 232, 211. Madison, J. T., Everett, G. A., and Kung, H.-K. (1967). JBC 242, 1318. Madison, J. T., Holley, R. W., Poucher, J. S., and Connett, P. H. (1967). BBA 145, Cantor, C. R. (1968). Biopo/ymers 6, 369. Burgers, P. M. J . , and Eckstein, F. (1979). Biochemistry 18, 450.

17. POLYNUCLEOTIDE PHOSPHORYLASE

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3. Equilibriiim PNPase directs either phosphorolysis of polynucleotides or polymerization of NDPs, depending on the reaction conditions and on the concentration of these two components in the reaction mixture. The mechanisms by which the enzyme drives these two reactions have been studied extensively with PNPase from M . futeus (82, 86) and from E . coli (85, 116), and detailed models have been proposed to explain the interrelations between various kinetic parameters that affect the dynamic equilibrium reached by the enzyme. The affinity of M . lirtrirs PNPase for either inorganic phosphate or oligonucleotide substrate is unaffected by the presence of either, and the initial rate of phosphorolysis depends linearly on the concentration of both. Oligonucleotides in which the 3'-OH group is blocked with a phosphate group are competitive inhibitors with respect to unblocked oligonucleotides, and noncompetitive with respect to inorganic phosphate. In contrast, the kinetics of phosphorolysis of polynucleotides shows that dADP is a competitive inhibitor with respect to both Pi and polynucleotide (86). Copolymerization of various NDPs occurs with M . lufeirs PNPase in a random fashion, indicating no special preference for any of the four common NDP substrates (f2.5). 4. Modified Sirbstrutes

Modifications of the NDP substrates serve to characterize the catalytic processes driven by the enzyme, the specificity of substrate recognition, and the properties of the active sites. Thus, blocking of the NDP at the 3' position yields a monovalent substrate, of which only one residue may be added to an oligonucleotide primer (126-128). It was shown that deoxynucleoside diphosphates are added to the 3' terminus of an oligonucleotide to a limited extent. The reason the polymerization of dADP cannot proceed readily seems to be due to the low affinity of the catalytic center (subsite I) to the DNA-like internucleotide linkage. When deoxyadenyl residues are added to the growing end of the chain, its a f h ity to the enzyme is lowered and the rate for further elongation is hence greatly reduced (I 13). Deoxyribonucleotides also act as inhibitors of PNPase. dADP inhibits competitively both the polymerization of ADP and the phosphorolysis of polynucleotides (86, 99), indicating that the 125. Seliger, H., and Knable, T. (1978). Nucleir Acids R r s . , Spec. Public. 4, S167. 126. Kaufmann, G., and Littauer, U. 2. (1970). EJB 12, 85. 127. Kaufmann, G., Fridkin, M., Zutra, A., and Littauer, U. 2. (1971). EJB 24, 4. 128. Bennett, G . N., Mackey, J. K . , Wiebers, J. L., and Gilham, P. T. (1973).Biochernistry 12, 3956.

534

U . Z. LITTAUER AND H.SOREQ

oligonucleotide primer covers the NDP binding subsite. However, when Mg2+is replaced by Mn2+,dADP is capable of copolymerizing with ADP (129). Other analogues, such as the periodate oxidation product of ADP, will block polymerization altogether (97) (see previous sections). PNPase displays a rather low specificity with regard to side chains on the purine or pyrimidine moieties (see previous sections), whereas it shows high specificity with respect to the number of phosphate groups on the nucleoside and the nature of the sugar moiety of the NDP substrate [cf. Ref. S ) ] . The polymerization parameters for various modified bases also serve to detect functional differences between PNPase from various strains of bacteria (130).

B. NUCLEOSIDE DIPHOSPHATE-P, EXCHANGE Two mechanisms were suggested for NDP-Pi exchange reaction: (1) The observed exchange reflects a reversible formation of a covalent, nucleoside monophosphate-enzyme complex, or (2) the apparent exchange is a result of combined polymerization and phosphorolysis reactions, occurring under approximate equilibrium conditions (8,39).The kinetic parameters of the exchange reaction appear to be similar to those of the polymerization reaction: It is preceded by a lag phase, activated by primers (8, 39), and occurs, to a limited extent, with deoxy NDPs, but only in the presence of oligonucleotide primers or NDPs (99, 113, 131). It was suggested that the use of dADP might facilitate isolation of the putative NMP-enzyme intermediate (113). However, no evidence for its formation could be obtained (131). Further support that the NDP-Pi exchange is the result of combined polymerization and phosphorolysis reactions is suggested from the arsenolysis of NDPs. Replacement of Pi by arsenate in the exchange reaction results in the arsenolysis of NDPs to nucleoside monophosphates. In the presence of primer-dependent PNPase from M. Iiiteus, arsenolysis of ADP, like its polymerization, is activated by oligonucleotides that have unesterified 3'-hydroxyl groups (116, 132). The kinetics of this reaction are consistent with the formation of a ternary complex between enzyme, oligonucleotide, and NDP. The formation of a new phosphodiester bond between the NDP and oligonucleotide and its subsequent arsenolysis is proposed for this reaction (132). A similar exchange reaction is catalyzed by a yeast ADP-sulfurylase, which does not show specificity for the sugar moiety, the nature of the NDP substrate, or 129. 130. 131. 132.

Chou, J. Y., and Singer, M. F. (1971). JBC 246, 7505. Swierkowski, M., and Shugar, D. (1969). Actu Biocliim. Polon. 16, 263. Chou, J. Y . , and Singer, M. F. (1971).JBC 246, 7486. Singer, M . F. (1963).JBC 238, 336.

17. POLYNUCLEOTIDE PHOSPHORYLASE

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the type of anhydride bond, and does not phosphorolyze polyribonucleotides. The mechanism by which the yeast enzyme catalyzes the exchange reaction appears to be a displacement of phosphate from NDP by different anions through formation of an intermediate AMP-enzyme complex (133). The kinetic parameters of the PNPase-directed exchange reaction were monitored by the appearance of an isotopic (I8O)shift in 31PNMR profile. yielded, during the exchange reaction, an L Y - P ( ' ~ O ~and ~~O a) Pi ('"04) /3-P(lH04),proving that bond cleavage occurs between the a-P and a-/3 bridge oxygen (134).

C. PHOSPHOROLYSIS In the presence of inorganic orthophosphate, PNPase acts as an exonuclease, releasing NDPs sequentially from the 3'-OH end of the polynucleotide substrate (135). PNPase readily phosphorolyzes single-stranded polynucleotides, but acts more slowly on multistranded structures (136, 137), or on polynucleotides with an extensive secondary structure, such as tRNA, rRNA (83, 137-/39), or mRNA [except the poly(A) tail, which is degraded rapidly; see Ref. 1/40)].The rate of phosphorolysis of RNA chains can be increased by raising the temperature of the reaction mixture (137, 140). The presence of a phosphate group at the 5' end does not prevent phosphorolysis. However, polyribonucleotides with a 3'-terminal phosphate group are not phosphorylyzed by the enzyme (8, 141). Dinucleotides, dinucleoside monophosphates, and, in some cases, trinucleotides are not substrates for phosphorolysis and these compounds accumulate as resistant end products (1 14, 120-122). PNPase phosphorolyzes short oligonucleotides (n 5 12) by a nonprocessive mechanism (82, 85, 112, 115, 142). In contrast, the enzyme tends to phosphorolyze long ( n I 133. Grunberg-Manago,M., Del Campillo-Campbell, A . , Dondon, L., and Michelson, A. M . (1966). BBA 123, 1. 134. Cohn, M . , and Hu, A. (1978). P N A S 75, 200. 135. Hilmoe, R . J . (1959). Ann. N . Y. Arnd. Sci. 81, 660. 136. Ochoa, S. (1957). ABB 69, 119. 137. Grunberg-Manago, M . (1959). JMB 1, 240. 138. Littauer, U. 2.. and Daniel, V. (1962). In "Acides Ribonucleiques et Polyphosphates," Colloq. Intern. du C.N.R.S., Strasbourg, p. 277. C.N.R.S., Paris. 139. Kimhi, Y. (1966). Doctoral Thesis, The Weizmann Institute of Science, Rehovot, Israel. 140. Soreq, H . , Nudel, U., Salomon, R., Revel, M . , and Littauer, U. 2. (1974). I M B 88, 233. 141. Singer, M. F., Heppel, L. A., Hilmoe, R. J., Ochoa, S . , and Mii, S. (1959). Ccm. Cancer Cotif. 3, 41. 142. Kaufmann, G . , Grosfeld, H . , and Littauer, U . 2. (1973). FEES Lerr. 31, 47.

536

U. Z. LITTAUER AND H. SOREQ

20) polynucleotides by a processive mechanism (i.e., the enzyme phosphorolyzes a single chain almost to completion before dissociating to initiate the phosphorolysis of another chain) (82, 83, 9G). The length of substrate at which the transition occurs between the two mechanisms depends on the sequence and the structure of the oligonucleotide (115, 142). Arsenate ions can replace inorganic phosphate in the degradation of polynucleotides by PNPase. The arsenolysis of polynucleotides liberates 5’-phosphorylarsenate nucleotides, which spontaneously hydrolyze to nucleoside monophosphates, arsenate, and H+ ions (143). Aminoacylated . tRNA chains can be phosphorolyzed by PNPase (126, 144). This phosphorolysis occurs by a similar mechanism to that observed with synthetic polynucleotides, as was shown for arsenolysis of valyl-tRNA, which yielded a valyl-adenosine monophosphate product (126). It has been shown that at 37” PNPase phosphorolyzes only part of tRNA molecules present in the reaction mixture, whereas the remaining chains appear to be completely intact (15). To phosphorolyze all the tRNA chains by the enzyme, the temperature of the reaction mixture has to be elevated over 45”, to permit a configurational change of the tRNA (83, 139). Under these conditions, PNPase phosphorolyzes tRNA in a processive mechanism and, similar to the degradation of synthetic polynucleotides, only NDPs and long substrate chains are present in the reaction mixture until the completion of the phosphorolysis reaction (83, 90). The configurational requirements that permit phosphorolysis of tRNA by PNPase are not related to the integrity of the anticodon loop, as tRNAP,h,e,reconstituted from split half molecules still retains the ability to undergo the change in conformation that permits phosphorolysis to occur (145). The transition between the two configurations of tRNA appears to involve large entropic changes, as shown for total unfractionatedE. coli tRNA (84) as well as for purified specific tRNA species (146n). The transition between the two configurational states appears to be initiated at a single “nucleation” center on the tRNA molecule (147, 148). The existence of multiple subsites for the interaction of PNPase with polynucleotides has been indicated from the various K, values that the enzyme displays with synthetic oligonucleotides of different lengths (1 14). This model was further substantiated by the comparative analysis of the 143. Singer, M. F., and O’Brien, B. M. (1963). JBC 238, 328. 144. Yot, P., Gueguen, P:,and Chapeville, F. (1968). FEBS Lett. 1, 156. 145. Beltchev, B., and Thang, M. N. (1970). FEBS Lett. 11, 55. 146. Beltchev, B . , Thang, M. N., and Portier, C. (1971). EJB 19, 194. 146a. Thang, M . N., Buckingham, R. H . , and Dondon, L. (1975). EJB 54, 93. 147. Danchin, A. (1972). FEES Lett. 19, 293. 148. Danchin, A,, and Thang, M . N. (1972). FEBS Lett. 19, 297.

17. POLYNUCLEOTIDE PHOSPHORYLASE

537

kinetic parameters for the PNPase-directed phosphorolysis of long polynucleotides, as compared with short oligonucleotide chains (cf. 15). It should be kept in mind that the enzyme molecule appears susceptible to conformational alterations, depending on the substrate present (86). Michaelis constants for short oligonucleotides are higher than those observed for long polynucleotide chains, the transition occurring at 40 2 n 2 10 (85). The detailed equilibrium constants for the phosphorolysis reaction catalyzed by PNPase under physiological conditions have been determined, and the rate of phosphorolysis was found to be sensitive to variations in free Mg2+but relatively insensitive to changes in pH (149).

D. “TRANSNUCLEOTIDATION” PNPase has been shown to catalyze the transfer of nucleoside phosphate moieties from a polynucleotide donor to a polynucleotide acceptor (141). The polynucleotide rearrangement arises from a combination of phosphorolytic and addition reactions of NDPs, catalyzed by trace amounts of inorganic phosphate contaminating the reaction mixture (150). Addition of a phosphate removal system consisting of calf spleen phosphorylase and nicotinamide riboside will block this ‘‘transnucleotidation” reaction. IV.

Attributed Physiological Functions

The apparent ubiquity of PNPase in microorganisms suggests an important role in cell physiology; however, an unequivocal demonstration of its biological function is still lacking. The function of the enzyme has been explored in E. coli mutant cells deficient in PNPase (151-IS), in toluenized cells (62, 156, 157), and in osmotically shocked cells (158). In spite of its widespread occurrence in bacteria, the enzyme is not indispensable to cell metabolism. Escherichia coli mutant cells with defective (152) or very low PNPase activity (155) show no difference in their growth rate at 37”, but grow somewhat more poorly at 45” than their revertants (159). These 149. 150. 151. 152. 153. 154. 155. 156. 157. 158. 159.

Liegel, J . , and Guynn, R. W. (1979). JBC 254, 1992. Sninsky, J . J . , Bennett, G. N., and Gilham, P. T. (1974). Nucleic Acids Res. 1, 1665. Reiner, A. M. (1%9). J B 97, 1431. Reiner, A. M. (1969). J B 97, 1437. Krishna, R. V., Rosen, L., and Apirion, D. (1973). Nature, New B i d . 242, 18. Kinscherf, T. G., Lee, Y.F., and Apirion, D. (1974). Nucleic Acids Res. 1, 1439. Portier, C. (1980). Molec. Gen. Genet. 178, 343. Levin, D. H., Thang, M. N., and Grunberg-Manago, M. (1963). BBA 76, 558. Deutscher, M. P. (1978). JBC 253, 5579. Raue, H. A . , and Cashel, M. (1974). BBA 340, 40. Krishna, R. V., and Apirion, D. (1973). IS 113, 1235.

538

U. Z. LITTAUER AND H. SOREQ

nonlethal mutations affect the structural gene for the a-chains of E. coli PNPase and map close to the argC locus (151, 155). It was suggested that PNPase participates in RNA metabolism (160, 158), and that in contrast to nucleases that liberate nucleoside monophosphates from RNA, PNPase conserves phosphate bond energy by releasing NDPs (160, 161). The liberated NDPs can later be reutilized for RNA synthesis or reduced to dNDP and incorporated after phosphorylation into DNA (160). The participation of PNPase in rRNA and mRNA metabolism has, therefore, been sought. It appears, however, that PNPase is not directly involved in the depolymerization of RNA in E. coli cells (162, 153). Examination of several PNPase deficient E . coli mutants suggests a possible role for PNPase as a salvage enzyme involved in rRNA or mRNA degradation in stressed cells starved for carbon at 49" (69, 163-166). In addition, analysis of PNPase mutants implies that the enzyme may participate in lcic mRNA degradation in heat-shocked cells (167). PNPase may be involved in the inactivation of extraneous eukaryotic mRNA. The expression of the catabolic dehydroquinase gene ( g a - 2 ) from Neurospora C Y Q S S ~is increased as much as 100-fold when cloned in E. cofi strains deficient in PNPase. These results suggest that there are inherent structural differences between prokaryotic and eukaryotic mRNAs (168). It has been suggested that PNPase may have a role in stabilization of mRNA chains by modifying their 3' ends. Comparison of the thermolabile PNPase mutant PR27 to its isogenic counterpart PR100 shows that at 37" or 45" the synthesis of /3-galactosidase proceeds at about the same rate. However, at 49" the functional haif-life of /3-galactosidase is shorter in the PNPase mutant cells (153,159). Several experiments suggest that PNPase could play a role in polyadenylation of mRNA (157, 169, 170). Possibly the poly(A) tail on E . coli mRNA would have a stabilizing function, as has 160. Sekiguchi, M., and Cohen, S. S . (1963).JBC 238, 349. 161. Tumerman, L., and Ric, S . (1977). "Applications of Calorimetry in Life Sciences," p. 97. Walter de Gruyter, Berlin and New York. 162. Chaney, S. G . , and Boyer, P. D. (1972);.J M B 64, 581. 163. Kinscherf, T. G . , and Apirion, D. (1975). Molec. G m . Genet. 139, 357. 164. Kaplan, R., and Apirion, D. (1975). JBC 250, 1854. 165. Kaplan, R., and Apirion, D. (1975). JBC 250, 3174. 166. Cohen, L., and Kaplan, R. (1977). J B 129, 651. 167. Har-El, R., Silberstein, A . , Kuhn, J., and Tal, M. (1979). Molec. Gen. Genet. 173, 135. 168. Hautala, J. A., Bassett, C. L., Giles. N. H., and Kushner, S . R. (1979). P N A S 76, 5774. 169. Wunderli, W., Hutter, R., Staehelin, M., and Wehrli, W. (1975). U B 58, 87. 170. Ramanarayanan, M., and Srinivasan, P. R. (1976).JBC 251, 6274.

17. POLYNUCLEOTIDE PHOSPHORYLASE

539

been suggested for some eukaryotic mRNA species (171,172).In crude extracts of T2L phage-infected E. coli cells, poly(A) synthesis from ATP arises from the combined action of PNPase and ATPase (169). Poyriboadenylate polymerase isolated from E. coii PR7 PNPase mutant will use either ATP or ADP as a substrate, although in this case ATPase appears as an integral part of the enzyme (170).Poly(A) synthesis has also been examined in toluenized E. coli cells. Mutant cells PR7 and PR13, deficient in PNPase, were unable to synthesize poly(A) (157),which is in contrast to the experiments with crude extracts (170).It should be noticed, however, that none of the above mutants are completely devoid of PNPase, as assayed by their phosphorolytic activity (15.2).The possibility that PNPase exists as a multienzyme complex with ATPase is suggested by analysis of the antigenic composition of the plasma membrane of S . pyrogenes (26). V.

Research Applications

A.

POLYNUCLEOTIDE SYNTHESIS

PNPase has been found to be a useful tool for the synthesis of polynucleotides with varied composition, both in the presence and in the absence of primer oligonucleotides. 1. Homopolymers

A large variety of homopolyribonucleotides have been prepared with the aid of PNPase. Because of the tendency of poly(G) to form multistranded helices, the polymerization of GDP proceeds to a very limited degree (3, 5, 106,108). These difficulties may be overcome with PNPase from E. coli by raising the temperature to 60" and by the replacement of Mg" by Mn2+(67,173).Poly(G) can also be synthesized at higher temperatures (70") with the aid of B. stearothe~mopliilus PNPase (17)or with PNPase from Thermirs thrrmophillrrs, in the presence of Mg2+(174).Various preparations of polyinosinic acid synthesized by PNPase differed in their secondary structure andor tertiary conformation. These differences resulted in varied reactivity with anti-poly(1)-antiserum, as well as in dif171. Nudel, U . , Soreq, H . , Littauer, U. Z . , Marbaix, G., Huez, G . , Leclercq, M., Hubert, E . , and Chantrenne, H. (1976). EJE 64, 115. 172. Littauer, U. Z . , and Soreq, H. (1982). Progr. Nirclric Acid Rrs. 27, 53. 173. Thang, M. N . , and Grunberg-Manago, M. (1968). "Methods in Enzymology," Vol. 12B, p. 522. 174. Kikuchi, Y., Hirai, K., Hishinuma, F., and Sakaguchi, K . (1977). BEA 476, 287.

540

U . Z. LITTAUER AND H. SOREQ

ferent abilities to induce the production of interferon in virus-infected cells (175). The basis for the differences with anti-poly(1) antiserum could be due to variability in the amount of hypoxanthine that is accessible to the antibody (176). PNPase also polymerizes modified NDPs, although at relatively slow rates. The range of NDP modification can be further extended by the use of Mn2+ as a cofactor, or with a matrix-bound enzyme (177). Thus, PNPase catalyzes the synthesis of polypseudouridylic acid (178-18O), poly-Zthiocytidylic acid (181, 182), poly-Cthiouridylic acid (183, l M ) , polyribothymidylic acid (185), poly-5-methyluridylic acid (186 ), poly-5ethyluridylic acid (187), polyfluorouridylic acid (188), poly-8-chloroadenylic acid (189), poly-8-oxyadenylic acid (I90), polyfluoroadenylic acid (191), poly(0 6-methyl or ethy1)guanylic acid, (192), and many other homopolymers. Fluorescent nucleotide analogues (lin-benzo-ADP and lin-IDP) have been prepared for use as dimensional probes of PNPase binding sites (193, 194). In contrast, some modifications render the modified nucleotide unsuitable for polymerization by PNPase. These modified NDPs, such as 5-acetyluracl NDP, may act as weak inhibitors of the enzyme (195). Chemical modifications of specific groups in NDPs were an aid in asses175. Stollar, B. D., DeClercq, E., Drocourt, J.-L., and Thang, M. N. (1978). HE 82, 339. 176. Inouye, H., Fuchs, S. , Sela, M., and Littauer, U. Z. (1971). BEA 240, 594. 177. Brentnall, H. J., and Hutchinson, D. W. (1972). Tetrahedron Lett. 25, 2595. 178. Sasse, L., Rabinowitz, M., and Goldberg, I. H. (1963). BBA 72, 353. 179. Pochon, F., Michelson, A. M., Grunberg-Manago, M., Cohn, W. E., and Dondon, L. (1964). BBA 80, 441. 180. Goldberg, I. H. (1968). “Methods in Enzymology,” Vol. 12B, p. 519. 181. Scheit, K. H., and Faerber, P. (1971). EJE 24,385. 182. Faerber, P., Scheit, K. H., and Sommer, H. (1972). EJE 27, 109. 183. Eckstein, F., and Scheit, K. H. (1971). I n “Procedures in Nucleic Acid Research” (G. L. Cantoni and D. B. Davies, eds.), Vol. 2, p. 665. Harper and Row, New York. 184. Fiser, I., Scheit, K. H., and Kuechler, E. (1977). EJB 74, 447. 185. Griffin, B. E., Todd, A., and Rich, A. (1958). PNAS 44, 1123. 186. Grunberg-Manago, M., and Michelson, A. M. (1964). EBA 80, 431. 187. Biala, E., Jones, A. S., and Walker, R. T. (1980). Tetrahedron 36, 155. 188. Grunberg-Manago, M., and Michelson, A. M. (1964). EBA 87, 593. 189. Tavale, S. S., and Sobell, H. M. (1970). JMB 48, 109. 190. Folayan, J. O., and Hutchinson, D. W. (1977). BEA 474, 329. 191. Broom, A. D., Amamath, V., Vince, R., and Brownell, J. (1979). EBA 563, 508. 192. Mehta, J. R., and Ludlum, D. B. (1976). Biochemistry 15, 4329. 193. Leonard, N. J., Scopes, D. I. C., VanDerLijn, P., and Barrio, J. R. (1978). Biochemistry 17, 3677. 194. Leonard, N. J., and Keyser, G. E. (1979). PNAS 76, 4262. 195. Jones. A. S., Stephenson, G. P., and Walker, R. T. (1979). Tetrahedron 35, 1125.

17. POLYN UCLEOTIDE PHOSPHORYLASE

54 1

sing the influence of these groups on the physical and chemical properties of polynucleotides (196). Thus, the role of the 2’-hydroxyl group in RNA conformation has been studied with the aid of 2‘-modified polynucleotides (197-200). Polymerized uridine-5‘-diphosphorothioateexhibits a certain extent of protection against nucleolytic degradation (20f). Poly-S4methyl-4-thiouridylic acid displays a specific emission spectra at 520 nm (202). 2-Azaadenosine and 2-azainosine diphosphates (203), as well as 2-methyl- and 2-ethylthioadenosine diphosphates (204) can be polymerized to their respective homopolymers, and the modifications do not prevent the formation of double-stranded complexes by the modified polymers. Poly(2’-deoxy-2’-fluorodenylicacid) and poly(2’-chloro-2’deoxyinosinic acid) have rather similar properties to those of poly(A), but differ from poly(dA) (205, 206). In addition, poly-5-methoxyuridylic acid stimulated the binding of Phe-tRNA to 70 S ribosomes, although it was inactive in directing poly(Phe) synthesis (207). In contrast, poly(Zfluoroadeny1ic acid) codes for the synthesis of polylysine (191). 2. Heteropolymers Polymerization of a mixture of NDPs that contain different bases results in random copolymers. Purine NDPs in which the purine and ribose are in the syn-conformation (189) are poor substrates for PNPase. However, as is the case for GDP, they may be incorporated into copolymers to a varying degree with the normal NDPs. Thus, ribopolynucleotides that contain 8-substituted purine nucleotides, such as 8-bromoadenosine, 8-oxyadenosine, 8-bromoguanosine, 8-oxyguanosine, and 8-dimethylaminoguanosine (208), as well as 1-methyl-6-thioguanosine(209) are synthesized by copolymerization of the modified NDPs with ADP or 196. Michelson, A . M., Massoulie, J . , and Guschlbauer, W. (1967). Progr. Nucleic Acid

Res. 6, 83.

197. Szer, W., and Shugar, D. (1966). J M B 17, 174. 198. Zmudzka, B . , Janion, C., and Shugar, D. (1969). BBRC 37, 895. 199. Zmudzka, B . , and Shugar, D. (1970). FEBS L e f t . 8, 52. 200. Torrence, P. F., Bobst, A . M . , Waters, J . A , , and Witkop, B. (1973). Biochemistry 11, 3962. 201. Eckstein, F., and Gindl, H. (1969). FEBS Lett. 2, 262. 202. Scheit, K . H . (1970). BBA 209, 445. 203. Fukui, T., Kakiuchi, N . , and Ikehara, M. (1978). BBA 520, 441. 204. Fukui, T., and Ikehara, M. (1979). BBA 562, 527. 205. Ikehara, M . . Fukui, T., and Kakiuchi, N . (1978). Nucleic Acids Res. 5, 1877. 206. Kakiuchi, N . , Fukui, T., and Ikehara, M. (1979). Nucleic Acids Res. 6, 2627. 207. Hillen, W., and Gassen, H. G. (1979). BBA 562, 207. 208. Ikehara, M . , Tazawa, I., and Fukui, T. (1969). Biochemistry 8, 736. 209. Amarnath, V., and Broom, A. D. (1977). BBA 479, 16.

542

U. Z. LITTAUER AND H. SOREQ

GDP. Copolymers that contain other base analogs, such as xanthosine, N’-methyluridine, N‘-acetylcytidine, and many others, have also been prepared, and serve to examine the role of rare and of “nonsense” bases in directing in virro protein synthesis (210, 21 I). 5’-Mercaptouridine 5’-diphosphate has been copolymerized with UDP and the resulting copolymer, after formation of double-stranded complex with poly(A), served as a potent inhibitor for DNA-dependent RNA polymerase (RNAdependent DNA nucleotidyltransferase, EC 2.7.7.7) (212). In contrast, polynucleotides that contain Oz- and 04-alkyluridine (213) or 2-thiocytidine (214) serve as templates for RNA polymerase activity. Heteropolymers that contain 2‘-O-methyladenylic acid and 2’-O-methylcytidylic acid have also been prepared (215, 2161, and dihydrouridine was more efficiently incorporated into heteroribopolymers than l-(fl-D-ribofuranosy1)-(a + P)5,6-methyleneuracyI (217). The stereochemistry of PNPasedirected internucleotide bond formation has been probed by polymeriza1-thiodiphosphate), which tion of the exoisomer of adenosine Y-04 undergoes inversion of its configuration into the endoisomer when copolymerized with UDP by PNPase (124). The fluorescent analog of adenosine, 1-N6-etheno-2-azaadenosine,has been incorporated into heteropolymers with ADP, UDP, or IDP and provides means for probing the structure of these polymers (218). The effect of spin-labeled copolymers on the reaction catalyzed by avian myoblastosis virus RNA-dependent DNA polymerase was studied by PNPase-directed copolymerization of 4-thiouridine and uridine, and it was shown that increasing amounts of potentially reactive thiol groups (or spin labels) enhance the inhibitory properties of the copolymers as compared to poly(U) (219). Simplified methods for the large-scale preparation of homooligonucleotides (220) and of heterooligonucleotides that contain modified nucleosides (221) have also been reported. A description of the various approaches utilized to 210. Michelson, A. M., and Grunberg-Manago, M. (1964). BBA 91, 92. 211. Michelson, A. M., and Pochon, F. (1966). BBA 114, 469. 212. Ho,Y.-K., Aradi, J., and Bardos, T. J . (1980). Nitcleic Acids Res. 8, 3175. 213. Singer, B . , Fraenkel-Conrat, H., and Kusmierek, J. T. (1978). PNAS 75, 1722. 214. Kroger, M . , and Singer, B. (1979). Biochemistry 18, 91. 215. Rottman, F., and Johnson, K. L. (1969). Biochemistry 8, 4354. 216. Simuth, J . , Strehlke, P., Niedballa, U., Vorbruggen, H., and Scheit, K. H. (1971). BBA 228, 654. 217. Torrence, P. F., and Witkop, B. (1972). Biochemistry 11, 1737. 218. Yip, K. F., and Tsou, K. C. (1979). Biopo/ymers 18, 1389. 219. Warwick, P.E., Hakam, A., Bobst, E. V., and Bobst, A. M. (1980). PNAS 77,4574. 220. Shum, B. W.-K., and Crothers, D. M. (1978). Nucleic Acids Res. 5, 2297. 221. Schetters, H., Gassen, H. G . , and Matthaei, H. (1972). BBA 272, 549.

17. POLYNUCLEOTIDE PHOSPHORYLASE

543

synthesize various building blocks for polynucleotide synthesis has been reviewed (222).

B . SYNTHESIS OF OLIGONUCLEOTIDES WITH A DEFINED SEQUENCE Under high salt concentrations, PNPase adds only a few nucleotide residues to the 3‘ end of a dinucleotide primer. This property of the enzyme served for the first preparations of oligonucleotides of defined sequence (223). The addition of one or two guanyl residues to oligonucleotide primers is achieved by incubation with PNPase from Thermu.~ thermophilrrs at 37”. At this relatively low temperature, poly(G) formation is inhibited (224). Monovalent addition of GMP residue to guanosine-free oligonucleotides, obtained by T1 ribonuclease digestion of RNA, can also be carried out by the simultaneous action of PNPase and T1 ribonuclease (224-226). Similarly, copolymers with a terminal pyrimidine residue are obtained by polymerization of a mixture of purine and pyrimidine NDPs with PNPase in the presence of pancreatic RNase (227). Two functional regions can be defined in the NDP monomers that serve as substrates for the polymerization reaction catalyzed by PNPase: The P-phosphate residue, which is eventually released as inorganic phosphate, and the free 3‘-hydroxyl group of the incoming NDP, which becomes the new accepting terminus (126). Certain modifications of the sugar moiety of the NDP substrate may convert it to a “monofunctional” substrate for PNPase. Such NDP derivatives, blocked in their 3’-hydroxyl function (probably due to steric hindrance), do not sustain de novo polymerization but are able to transfer one nucleotidyl residue to an oligonucleotide initiator, thus serving as chain terminators. The blocking group can be subsequently chemically removed from the oligonucleotide product, permitting a succession of single addition reactions to be carried out. This procedure has been utilized for the stepwise synthesis of polyribonucleotides of defined sequence (126, 228). 222. Seliger, H . , Haas, B . , Holupirek, M., Knaeble, T.,Todling, G . , and Philipp, M . (1980). N d r i c . A t k f s l i c ? . ~ .. Syrnp. S r r . N O . 7. 191 . 223. Thach, R. E. (1966). Zn “Procedures in Nucleic Acid Research” (G. L. Cantoni and D. R. Davies, eds.), p. 520. Harper and Row, New York. 224. Kikuchi, Y . , Hirai, K . , and Sakaguchi, K. (1979). J B 86, 1427. 225. Szeto, K. S . , and Soll, D. (1974). Nucleic Acids Res. 1, 171. 226. Kikuchi, Y., and Sakaguchi, K. (1978). Nircleic-Acids Res. 5, 591. 227. Saunders, C . A., Sogin, S. J . , and Halvorson, H. 0. (1979). A B 95, 171. 228. Kaufmann, G . , Zutra, A., and Littauer, U. Z. (1971). Isr. J . Chem. 9, 44BC.

544

U. Z. LITTAUER AND H. SOREQ

NDPs containing a variety of blocking groups have been employed for the monovalent addition of a single nucleoside residue to a given oligonucleotide primer (229).These include the corresponding 2’(3’)-O-isovaleryl (127, 230), and 2’(3‘)-O-a-methoxyethyl(128, 150, 231 -234) diphosphates that were added to trinucleotide primers with a free 3’-OH group. After removal of the protecting groups by treatment with weak alkali (isovaleryl) or acid (methoxyethyl), the products can serve as acceptors for a second single-addition reaction. Oligonucleotides of defined sequence of four to seven residues have been synthesized by these methods (127, 228, 231). NDPs that contain 2‘(3’)-dihydrocinnamoyl (235) and the photolabile 2’-0-(0 -nitrobenzyl) groups have been utilized for the monoaddition to tri- and tetranucleotide primers (236, 237). The monoaddition reaction is accompanied by a limited rearrangement of the initiator oligonucleotide ( l 2 7 ) , which can be circumvented by coupling the reaction with an enzyme system that utilizes inorganic phosphate either present or formed in the reaction mixture (150, 238). Combination of these and other reactions, such as the use of T4 RNA ligase to ligate the synthesized oligonucleotides, permits the synthesis of oligoribonucleotides of defined sequence of appreciable length (226,239,240).NDPs in which the C-2’-C-3’ bond has been cleaved (ox-red nucleosides) by periodate oxidation followed by borohydride reduction may serve as monovalent terminators of PNPase-catalyzed polymerization, and can also be used for radioactive labeling of the 3‘ termini of polyribonucleotides (232). 229. Kossel, H., and Seliger, H. (1975). In “Recent Advances in Polynucleotide Synthesis” (W. Herz, H. Grisebach, and G. W. Kirby, eds.), p. 467. Springer-Verlag, Berlin and New York. 230. Walker, G. C., and Uhlenbeck, 0. C. (1975). Biorhernistry 14, 817. 231. Mackey, J . K., and Gilham, P. T. (1971). Natrire (London) 233, 551. 232. Hawley, D. M., Sninsky, J. J., Bennett, G. N., and Gilham, P. T. (1978). Biochemisrp 17, 2082. 233. The Nucleic Acid Synthesis Group, Shanghai (1979). Aria Biorhirn. Biophys. Sin. 11, 290. 234. Sninsky, J. J., Hawley, D. M., and Bennett, G. N. (1975). FP 34, 702. 235. Kikuchi, Y., Hirai, K., and Sakaguchi, K. (1975). J B 77, 469. 236. Ikehara, M., Tanaka, S., Fukui, T., and Ohtsuka, E. (1976). Nucleir Acids Res. 3, 3203. 237. Ohtsuka, E., Tanaka, S., Hayashi, M., and Ikehara, M. (1979). BBA 565, 192. 238. Kikuchi, Y., Someno, K., and Sakaguchi, K. (1977). Agr. Biol. Chem. 41, 1531. 239. Kaufmann, G . , and Littauer, U. Z. (1974). PNAS 71, 3741. 240. Gumport, R. I. and Uhlenbeck, 0. C. In “Gene Amplification and Analysis” (J. G. Chirikjian and T. S. Papas, eds.), Vol. 11, in press. Elsevier North Holland, New York.

17. POLYNUCLEOTIDE PHOSPHORYLASE

c.

POLYMERIZATION

OF

545

DEOXYRIBONUCLEOTIDES

PNPase is unable to phosphoroloyze DNA (8). However, the enzyme can direct the reversible addition of a single deoxynucleotidyl residue to ribooligonucleotide primers. Further addition of deoxynucleotide residues to the resulting product is very difficult (99, 113, 131, 241, 242). PNPase does not readily catalyze the de nuvo synthesis of (dA), chains, probably because it is a poor substrate for chain initiation (113, 131,243). However, in the presence of Mn2+, E. culi PNPase catalyzes the transfer of deoxyribonucleotide residues from dNDPs to the 3’ OH end of an oligodeoxyribonucleotide primer having a minimal length of three nucleoside residues. This allows the synthesis, by repeated addition of single residues, of oligodeoxyribonucleotidesof defined sequence, although the overall yield is rather poor. The kinetics of the addition reactions differ for various deoxyribonucleoside 5’-diphosphates and for different primers (244-248). The limited addition reaction displayed with deoxyribonucleoside diphosphates contrasts with the extended polymerization that has been observed for a number of dNDP derivatives that contain substituents at the C-2’ position (198-200, 217, 242, 243, 249-254). PNPase also adds to oligodeoxynucleotide primers modified deoxynucleoside diphosphates, such as 5-methyldeoxycytidine, N4-hydroxydeoxycytidine, and deoxyuridine. Some modifications, such as 5-mercurideoxyuridine, prevent the addition of the modified nucleoside base to deoxyribooligonucleotide primer by PNPase (255).

241. Feix, G . (1972). BBRC 46, 2141. 242. Batey, I . L., and Gilham, P. T. (1974). Biochemistry 13, 5395. 243. Rottman, F., and Heinlein, K. (1968). Biochemistry 7, 2634. 244. Gillam, S., Rottman, F., Jahnke, P., and Smith, M . (1977). P N A S 74, 96. 245. Gillam, S . , Jahnke, P., and Smith, M. (1978). JBC 253, 2532. 246. Gillam, S . , and Smith, M . (1980). “Methods in Enzymology,” Vol. 65, p. 687. 247. Wu, R., Bahl, C. P., and Narang, S. A. (1978). Progr. Nucleic Acid Res. 21, 101. 248. Trip, E. M., and Smith, M. (1978). Nucleic Acids R e s . 5, 1529. 249. Janik, B . , Kotick, M. P., Kreiser, T. H., Reverman, L. F., Sommer, R. G . , and Wilson, D. P. (1972). BBRC 46, 1153. 250. Hobbs, J., Sternbach, H . , and Eckstein, F. (1971). FEBS Lett. 15, 345. 251. Hobbs, J . , Sternbach, H . , and Eckstein, F. (1972). BBRC 46, 1509. 252. Hobbs, J . , Sternbach, H . , Sprinzl, M . , and Eckstein, F. (1972). Biochemistry 11, 4336. 253. Khurshid, M., Khan, A , , and Rottman, F. M. (1972). FEBS Lett. 28, 25. 254. Tazawa, I . , Tazawa, S . , Alderfer, J. L., and Ts’o, P. 0. P. (1972). Biochemistry 11, 493 1. 255. Trip, E. M . , and Smith, M. (1978). Nircleic Acids R e s . 5, 1539.

546 D.

U . Z. LITTAUER AND H. SOREQ

CONJUGATION T O INSOLUBLE MATRIX

PNPase from both E. coli (42,117,256-259)and M . Iuteus (256)has been bound to a variety of insoluble matrices, such as cellulose nitrate filters (260),cellulose beads (261),mercerized cellulose (256),Sepharose 4B (42, 47, 228, 256), hydrazide agarose (257), diazotized p-aminobenzenesulfonylethyl (ABSE) agarose, ABSE-Sephadex G-200, and ABSE-cellulose (258, 259). Cellulose-bound PNPase can polymerize NDPs under pH conditions at which phosphorolysis is negligible (256). The insoluble PNPase has therefore been used to improve the yield of the polymerization reactions, especially those that involve atypical bases and are difficult to carry out, such as poly 8-chloroadenylic acid (177) and poly(1) chains (257). Cellulose- and Sepharose-bound PNPase phosphorolyze polynucleotide chains at a slower rate than that of the soluble enzyme (42, 228, 260), and display K , values for long polynucleotides that are higher by two orders of magnitude than those measured for the soluble enzyme (117). The phosphorolysis of long RNA molecules by Sepharose-bound PNPase involves three active subunits, as has been titrated by the removal of poly(A) tails from globin mRNA (42). Unlike the soluble enzyme, bound PNPase phosphorolyzes polynucleotides by a nonprocessive mechanism, although the kinetic parameters of the phosphorlysis of short oligonucleotides are unaltered ( 1 17). The insoluble PNPase has several advantages over the soluble enzyme, both for analytical and preparative purposes. The same enzyme preparation can be recycled multiple times (42,258,259),and the separation of the reaction products from the enzyme is greatly simplified. Thus, insoluble PNPase has been used for the enzymatic synthesis of polynucleotides ( 1 17, 258, 259) as well as for controlled phosphorolysis of mRNA (42), of viral RNAs (262, 2631, and of whole TMV viral particles, in which the 256. Smith, J . C . , Stratford, I. J . , Hutchinson, D. W., and Brentnall, H. J. (1973). FEBS

Lett. 30, 246.

257. Bachner, L., De Clercq, E., and Thang, M. N . (1975). BBRC 63, 476. 258. Yang, K.-Y., Liu, N.-J., and Lin, Y. (1979). Acttr Biochim. Biophys. Sin. 11, 87. 259. Yang, K.-Y., Liu, N.-J., and Lin, Y. (1979). Acra Biochim. Biophys. Sin. 11, 104. 260. Thang, M. N . , Graffe, M., and Grunberg-Manago, M. (1968). BBRC 31, 1 . 261. Hoffman, C. H . , Harris, E., Chodroff, S., Michelson, S., Rothrock, J. W., Peterson, E., and Reuter, W. (1970). BBRC 41, 710. 262. Salomon, R . , Sela, I . , Soreq, H., Giveon, D., and Littauer, U. Z. (1976). Virology 71,74. 263. Salomon, R . , Bar-Joseph, M., Soreq, H . , Gozes, I., and Littauer, U. Z. (1978). Virology 90, 288.

17. POLYNUCLEOTIDE PHOSPHORYLASE

547

3‘-terminal nucleotides are vulnerable to the nucleolytic attack by PNPase even in the presence of the viral protein coat (264). E.

RADIOLABELED NUCLEOTIDES AND FINGERPRINTING OF OLIGONUCLEOTIDES S Y N T H E S I S OF

PNPase has been used to synthesize radiolabeled polyribonucleotides from NDP monomers (265,266). It has also been used for sequence analysis of short oligoribonucleotides. These are phosphorolyzed by PNPase starting from the 3’ end in a stepwise fashion, and by a nonprocessive mechanism (82, 85, 112, 1/.5), to yield a mixture of NDPs and a limit oligonucleotide that cannot further be degraded by the enzyme. One may use labeled oligonucleotides or include [32P]orthophosphatein the reaction mixture. By following the order in which the released P-labeled NDPs appear during the phosphorolysis of a given oligonucleotide, it is possible to determine the nucleotide sequence from the 3‘ end up to 2-3 residues from the 5’ terminus (142). This scheme served to develop a method for sequence analysis of short oligonucleotides. PNPase-directed labeling of nucleolytic cleavage-oligonucleotides, aids in the fingerprint analysis of RNA sequences. RNA fragments derived by T1 RNase are dephosphorylated with bacterial alkaline phosphatase to yield oligonucleotides with free 3‘-hydroxyl groups, which may in turn serve as primers for polymerization by PNPase. In the presence of as a substrate and T1 ribonuclease, only a single a-labeled [CX-~~PIGDP GMP is added to the pancreatic RNase-derived fragments (225). Several procedures that use PNPase have been developed for the labeling of NDPs and NTPs at their @-position. One of these utilizes the exchange reaction catalyzed by PNPase between [32P]inorganicphosphate and the @-phosphatemoiety of a given NDP. The [@-32P]NDP obtained can then be phosphorylated to generate the [P-32P]NTPderivative (267-270). Another technique that yields [@-32P]NTP with a very high specific activity, exploits the phosphorolysis properties of PNPase. According to this 264. Littauer, U . Z . , Soreq, H., and Cornelis, P. (1980). In “Enzyme Regulation and Mechanism of Action” (P. Mildner and B. Ries, eds.), FEBS, Vol. 60,p. 233. Pergamon, New York. 265. Leder, P., Singer, M. F., and Brimacombe, R. L. C. (1%5). Biochemistry 4, 1561. 266. Singer, M. F., Hilrnoe, R . J . , and Heppel, L. A. (1960). JBC 235, 751. 267. Littauer, U. Z . , Kimhi, Y., and Avron, M. (1964). AB 9, 85. 268. Gilboa, E., Soreq, H . , and Aviv, H. (1977). EJB 77, 393. 269. Vennstrom, B . , Pettersson, U . , and Philipson, L. (1978). Nircleic Acids Res. 5, 205. 270. Eliasson, R., and Reichard, P. (1978). JBC 253, 7469.

548

U. 2.LITTAUER AND H. SOREQ

procedure, a polyribonucleotide of choice is phosphorolyzed by PNPase in the presence of carrier-free 32P-inorganicphosphate. The resulting [p32P]NDPproduct is then phosphorylated by pyruvate kinase, which drives the reaction to completion (271, 272). [p-32P]Purinetriphosphates prepared by this method serve as useful precursors in studying the initiation of eukaryotic mRNA (268) and their 5'-terminal caps (273), as well as in studying the initiator RNA of short nascent DNA chains (Okazaki pieces) (270).

F. SYNCHRONOUS PHOSPHOROLYSIS AS A N ANALYTICAL TOOL 1. Removal of Poly(A) Tracts from mRNA

The 3'-exonucleolytic activity of PNPase has been used for &heanalysis of the size and composition of the 3'-terminal sequence of RNA molecules (140, 171, 172, 264, 274-278). The analysis is based on the property of the enzyme to phosphorolyze long polynucleotides by a processive mechanism. The use of molar excess of PNPase over the substrate establishes a synchronous mode of phosphorolysis, in which NDP molecules are sequentially released from the 3' terminus of the RNA chains. In order to follow the course of phosphorolysis, [32Plorthophosphateis included in the reaction mixture and the released &labeled NDPs are analyzed by DEAE-cellulose paper chromatography (140, 142) or by PEI-cellulose ascending thin-layer chromatography (27.5, 279). The size of the shortened RNA molecules is then determined by gel electrophoresis on polyacrylamide-agarose composite gels (140, 263) or in gels under denaturing conditions (276, 278). In some cases 32P-labeledor 13H]uridinelabeled RNA was included in the phosphorolysis reaction and the released NDPs are labeled accordingly (276, 278, 280). 271. Leung, K.-L., and Yamazaki, H. (1977). Can. J . Eiochem. 55,223. 272. Kaufmann, G., Choder, M . , and Groner, Y. (1980). AE 109, 198. 273. Groner, Y., Gilboa, E., and Aviv, H. (1978). Biochemistry 17, 977. 274. Littauer, U. Z., Salomon, R., Soreq, H., Fleischer, G . , and Sela, I. (1975). I n "Organization and Expression of the Viral Genome. Molecular Interaction in Genetic Translation" (F.Chapeville and M. GrunbergManago, eds.), Vol. 39, p. 133. Roc. 10th FEBS Meeting, Paris. 275. Vournakis, J. N., Efstratiadis, A., and Kafatos, F. C. (1975). PNAS 72, 2959. 276. Grosfeld, H . , Soreq, H . , and Littauer, U. Z. (1977). Nucleic Acids Res. 4, 2109. 277. Kaempfer, R . , Hollender, R., Soreq, H., and Nudel, U. (1979). EJB 94, 591. 278. Soreq, H., Sagar, A. D., and Sehgal, P. B. (1981). PNAS 78, 1741. 279. Deshpande, A. K., Chatterjee, B., and Roy, A. K . (1979). JEC 254, 8937. 280. Sehgal, P. B., Soreq, H., and Tamm, I. (1978). PNAS 75, 5030.

17. POLYNUCLEOTIDE PHOSPHORYLASE

549

At 0" the poly(A) tails of mRNA molecules are readily phosphorolyzed while the rest of the RNA chains remain intact (140, 171). The rate of poly(A) phosphorolysis varies with the ionic strength, and ranges between 7.5 nucleotides per chain per minute at a 1.0 M NaCl concentration and 75.0 nucleotides per chain per minute at an ionic strength of 0.15 M (140, 264). The calculated rate of phosphorolysis is based on the assumption that all RNA chains are bound to enzyme molecules and phosphorolyzed synchronously at the same rate. Analysis of a heterogeneous RNA population therefore yields an average rate measurement. This implies that the measured rate of phosphorolysis may be underestimated in cases where not all the chains possess poly(A) tails (264). The length of the phosphorolyzed poly(A) tail has been estimated by comparative gel electrophoresis of native and deadenylated mRNA and by determination of the number of moles of ADP that are liberated per mole of RNA (140, 276). Using these methods, it has been established that the average size of the poly(A) tail for different preparations of rabbit globin mRNA is between 120-150 residues (140, 277). Similar experiments with the rat liver mRNA for a-2p-globulin revealed a variety of lengths for its poly(A) tails, ranging between 40 to 175 residues (279). 2. Phosphorolysis qf 3 ' Seyirences from R N A As mentioned above, at 0" only poly(A) tails are phosphorolyzed, possibly due to the difference in secondary structure between homopolynucleotides and heteropolynucleotides. Raising the temperature of incubation to 37" allows the phosphorolysis of heterogeneous sequences from RNA populations. Even at this elevated temperature and at low salt concentration, the rate of phosphorolysis varies greatly among the RNA species tested. Thus, after the removal of the poly(A) region from globin mRNA at O", the rest of the RNA chains are phosphorolyzed synchronously at 37" at an average rate of 9 nucleotides per chain per minute (227). A similar rate of phosphorolysis is observed with the larger (1.4 X lo6MW) RNA from Carnation mottle virus (263). TMV RNA, in contrast, is phosphorolyzed at a much slower rate of 3.5 nucleotides per chain per minute (274). This may result from the more compact conformation of the tRNAlike structure at the 3' end of the TMV RNA, that is specifically aminoacylated with histidine (262). In vivo protection of RNA sequences at their 3' end may also be indicated from the fact that rho protein, an RNA synthesis termination factor fromE. coli, binds tightly to poly(C) or poly(U) and prevents their degradation by PNPase (281). 281. Galluppi, G . R.,and Richardson, J . P. (1980). J M B 138, 513.

550

G.

U. Z. LITTAUER A N D H. SOREQ PROBE FOR T H E REGULATORY FUNCTION OF T H E 3'-OH REGIONOF RNA

1. Regulatory Role of Poly(A) fvom Vurious mRNA Species

Synchronous phosphorolysis of RNA by PNPase has been used to examine the role of the 3' nontranslatable regions of RNA chains (cf. 172). Highly purifiedE. coli polynucleotide phosphorylase (42)was used to phosphorolyze the poly(A) tracts of rabbit globin mRNA under conditions in which poly(A) is removed but the rest of the molecule remains intact (140, 264). The deadenylated globin mRNA is translated in vitro as efficiently as native mRNA for a short while. Upon longer periods of incubation, the rate of protein synthesis decreased more rapidly with the deadenylated mRNA than with the native mRNA (140), suggesting that the presence of the poly(A) sequence may stabilize the functional activity of mRNA molecules in vitro. This stabilization is not limited to mRNA molecules. Addition of poly(A) segments to E. coli 5 S RNA, which is also carried out with the aid of PNPase, increased the stability of the 5 S RNA against endonucleolytic attacks (282). However, the interaction between globin mRNA and the initiation factor that binds methionyl tRNAfMe'remained unchanged when phosphorolysis was used to remove the poly(A) tail or even the 90 nucleotides adjacent to it in the untranslated 3' sequence (277). In contrast, deadenylation of ovalbumin mRNA by PNPase was reported to reduce the initiation process (283). It is not clear, however, whether o r not other regions of the mRNA were altered as well during the deadenylation procedure, reported for ovalbumin mRNA. The removal of the poly(A) region clearly decreases the functional and physical stability of globin mRNA in microinjected Xenopus oocytes. Following an equilibration period, the native poly(A)-containing globin mRNA remains fully active for at least 72 hr (284), and so does globin mRNA species from which their poly(A) tails are shortened down to 32 adenylate residues (171). Poly(A)-free globin mRNA and a globin mRNA population that contains an average length of 16 adenylate residues per chain showed a rate of decay with a of about 6 hr. Thus, the poly(A) region must contain a minimal number of about 30 adenylate residues to ensure its protective function ( 1 71). The stabilizing role of poly(A) on mRNA is not a general phenomenon (172). Thus, the physical and functional stability of human interferon 282. 283. 284. trenne,

Hieter, P. A . , LeGendre, S. M . , and Levy, C. C. (1976). JBC 251, 3287. Doel, M . T., and Carey, N . H. (1976). Cell 8, 51. Marbaix, G., Huez, G., Burny, A., Cleuter, Y., Hubert, E . , Leclercq, M., ChanH., Sores, H., Nudel, U . , and Littauer, U. Z. (1975). P N A S 72, 3065.

17. POLYNUCLEOTlDE PHOSPHORYLASE

55 1

mRNA species in microinjected Xenoprrs oocytes is not affected by the removal of poly(A) tails from their 3' termini with PNPase (278, 280). Since poly(A) tails exist on most species of mRNA, it appears that the biological role of poly(A) other than as a stabilizing element remains to be revealed.

2. Role (d3' Termini in tRNA and in rRNA Replacement of the 3'-terminal adenosine moiety of tRNA with 2'- and 3'-deoxyadenosine afforded tRNA species useful in defining the nature of the partial reactions which comprise protein biosynthesis. Thus, incubation of an enzymatically abbreviated tRNA (tRNA-C-COH) with 2'deoxy-3'-O-~-phenylalanyladenosine and PNPase yielded tRNA terminating with the corresponding aminoacylated deoxynucleoside. The yield of this product is increased by including 20% methanol in the reaction mixture (285). Processive removal of 160 nucleotides from the 3' end of E. coli 16 S rRNA was found to have little if any effect on the ability of the phosphorolyzed rRNA to be reconstituted into 30 S ribosomal subunits, which contain all of the native ribosomal proteins and bind formylmethionyltRNA with equal efficiency t o native 30 S subunits, but have low capacity to direct protein synthesis (286). 3. Role of 3' Sequences in Viral R N A PNPase has been extensively employed to reveal the role of 3' sequences in numerous polyadenylated as well as poly(A)-deficient viral RNA species. The poly(A) tail of mRNA from Sendai virus (287) and from measles virus (288) have been shown to be nonexposed to exonucleolytic attack by PNPase. Deadenylation of poliovirus RNA abolishes its infectivity, as a result of the inability of the RNA to serve as a template for the viral replicating enzyme (289). Several viral RNA and viral mRNA species contain mixed nucleotide sequences at the 3' end, rather than a poly(A) tail. The 3' end of TMV RNA, which is devoid of a poly(A) tail, is essential for its infectivity. It has been observed that synchronous phosphorolysis of about 5 nucleoside residues per chain completely abolished the infectivity of the phosphorolyzed TMV RNA (262). Even within the viral particles, the same 285. Chinault. A. C . , Kozarich, J. W., Hecht, S. M . , Schmidt, F. J . , and Bock, R. M. (1977). Biochernb/ry 16, 756. 286. Zagorska, L., Szkopinska, A., Klita, S., and Szafranski, P. (1980). BBRC 95, 1152. 287. Marx, P. A., Jr., F'ridgen, C., and Kingsbury, D. W. (1975). J . Gen. Virol. 27, 247. 288. Hall, T. C. (1979). fntern. R e v . Cytol. 60, 1. 289. Dasgupta, A , , Zabel, P., and Baltimore, D. (1980). Cell 19, 423.

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3'-terminal nucleotides of TMV RNA appear to be vulnerable to exonucleolytic attack, and their removal by immobilized PNPase destroys their infectivity (264 ). A plant virus of a different architectural design is the Carnation mottle virus (CarMV), consisting of round particles with no vulnerable termini. When translated in cell-free extracts, CarMV RNA operates as a polycystronic message, which induces the synthesis of three distinct polypeptides of molecular weights of 77,000, 38,000, and 30,000. The 38,000 polypeptide is the subunit of the viral coat protein. In contradistinction with TMV RNA, the infectivity and the translational activity of CarMV RNA chains is gradually reduced following the removal of 3'-end sequences with the aid of PNPase. The rate of decrease of the infectivity is faster than the ability to sustain in v i m translation of the viral coat protein. Moreover, the reduction in the rate of synthesis of the 77,000 product is even faster than loss of infectivity. These observations imply that this unidentified large polypeptide, but not the viral coat protein, may be essential for infection and that the translation of CarMV RNA into this protein is highly dependent upon the intactness of the vulnerable 3' end (263). 4. Role of 3'-Noncoding Sequences in mRNA

The coding regions in all known cases of mRNAs, whether polyadenylated or not, are followed by 3'-nontranslated heteropolymeric sequences, which differ in length for individual mRNA species. It appears that phosphorolysis of the entire 3'-noncoding region, including the AAUAAA hexanucleotide transcript, does not abolish the translational efficiency of rabbit globin mRNA (277,264).Similar conclusions were drawn when the entire 3'-noncoding sequence was deleted from globin mRNA by doublestranded nuclease, following its hybridization to a cloned cDNA probe (290). Moreover, the 3'-noncoding region does not participate in the formation of the initiation complex, since the interaction between globin mRNA and the initiation factor that binds methionyl tRNAfMetremained unchanged when the entire 3'-noncoding region was removed by phosphorolysis with PNPase (277). The phosphorolytic removal of the poly(A) tails and the entire 3'noncoding regions from interferon mRNA species with the aid of PNPase does not significantly alter the translational efficiency or stability of these molecules when microinjected into Xenopus oocytes (278).Therefore, the AAUAAA hexanucleotide, which is included in the deleted region, and 290. Kronenberg, H. M . , Roberts, B. E., and Efstratiadis, A. (1979). Nucleic Acids Res. 6 , 153.

553

17. POLYNUCLEOTIDE PHOSPHORYLASE

the whole of the 3'-noncoding region, do not appear to contribute to the regulation of interferon mRNA stability in the Xenoprrs system.

H. PNPAsE-DIRECTED LABELING OF POLY N uc L EOTI DES

THE

3'-OH ENDOF

Primer-dependent PNPase has been used to add poly(C) sequences to the 3' terminus of RNA from potato spindle tuber viroid, and the resulting RNA served as template for QP replicase. The poly(G) sequence at the 5' end of the product provides a potential means to separate template from product and to study the properties of both RNA chains (291 ). Polyadenylation of viral RNA species makes them substrates for reverse transcription. The resulting labeled cDNA can then be sequenced, as was carried out with RNAs purified from preparations of vesicular stomatitis virus (292). The ability of PNPase to add poly(A) tails to the 3'-OH end of RNAs (tRNA, 5 S RNA or poly(A)-deficient mRNA) was also utilized for gene mapping. The poly(A) tailing is accomplished by use of a 2- to 3-fold excess of PNPase over RNA and 20-200 I.LMADP. Under these conditions 10-2096 of the RNA molecules acquire a poly(A) tail of about 60-400 residues long. The poly(A)-containing RNA is separated from the nonreacted RNA by oligo(dT)-cellulose chromatography. The in v i m polyadenylated RNA is then hybridized with a linear duplex DNA to which poly(dT) tails, or poly(dBrU) tails have been added with terminal deoxynucleotidyltransferase. The poly(dT) pairs with the poly(A) on the RNA and is readily recognized in the electron microscope (293).

291. Owens, R. A., and Diener, T. 0. (1977). Virology 79, 109. 292. Rowlands, D. J. (1979). PNAS 76, 4793. 293. Engel, J. D., and Davidson, N. (1978). Biochemistry 17, 3883.