Mammalian Nucleolytic Enzymes and Their Localization

Mammalian Nucleolytic Enzymes and Their Localization DAVIDSHUGAR AND HALINASIERAKOWSKA Institute of Biochemistry and Biophysics, Academy of Sciences, and Department of Biophysics, University of Warsaw, Warsaw, Poland

I. Introduction . . . . . . . . . 11. Types of Nucleolytic Enzymes . . . . A. Endonucleases . . . . . . . . B. Exonucleases . . . . . . . . C. Cyclic Nucleotide Phosphodiesterases . . 111. Methods of Assay . . . . . . . . A. Endonucleases . . . . . . . . B. Natural Inhibitors . . . . . . . C. Exonucleases . . . . . . . . D. Cyclic Nucleotide Phosphodiesterases . . IV. Substrate Preparations . . . . . . . V. Cellular Fractionation . . . . . . . VI. Histochemical Methods . . . . . . VII. Cytochemical Procedures . . . . . . A. Immunofluorescence Techniques . . . B. Precipitate-Forming Techniques . . . C. Comparison of Cytochemical and Fractionation VIII. Nuclealytic Enzymes in Pathological States . Nucleases in Tumors . . . . . . . IX. Possible Functions of Nucleolytic Enzymes . Addendum . . . . . . . . . . References . . . . . . . . Note Added in Proof . . . . . . .

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1. Introduction A multitude of enzymes of varying degrees of specificity exhibit hydrolytic activity against RNA, DNA, and nucleoside cyclic phosphates. Relatively little is known about the biological function of these enzymes and, in particular, of their intracellular localization, both of these being, of course, interrelated. Even if we discount the difficulties normally encountered in investigations on enzyme localization, those involving nucleolyt.ic enzymes have been particularly arduous because of the wide range of enzymes with, frequently, overlapping specificities. 369

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The following brief review makes no claim to provide any clear-cut solutions to these problems. Its preparation was prompted rather by the fact that, during the past five years, the preparation of some specific substrates, and the development of methods for their application to cell fractionation techniques and histo- and cytochemical procedures, has a t least laid a reasonably solid foundation for future work. Furthermore, it is felt that no apology is required for the inclusion of brief references to some studies now known to be of questionable validity, but the principles of which may nonetheless prove of value in an important field still undergoing birth pangs.

II. Types of Nucleolytic Enzymes Attempts a t the direct intracellular localization of nucleolytic enzymes have been confined largely t o the differentiated materials of higher organisms. The following outline is therefore limited to mammalian enzymes, although reference is made below to some bacterial enzymes (Section IX) in connection with a discussion of their function. Only essential properties are listed, such as substrate specificity, type of attack, and nature of products; additional aspects are included when they appear to be of value in distinguishing between different enzymes in localization studies. Further details may be found in several review articles (1-9).

A. Endonucleases 1. RIBONUCLEODEPOLYMERASES

Pancreatic R N m e (EC 2.7.7.16). This enzyme is most abundant in the pancreas (10, I l ) , but similar activity is found in other tissues and body fluids (12-16). It is stable to acid and heat, and is optimally active a t pH 7-8. Its activity may be “masked” by natural inhibitors in mammalian tissues (16-20). Its action is biphasic and involves first the rapid hydrolysis of the internucleotide linkage between a pyrimidine nucleoside 3’-phosphate and the adjacent nucleoside to give a pyrimidine nucleoside 2’:3’-cyclic phosphate; this is followed by the much slower hydrolysis of the cyclic phosphate ring to give a pyrimidine nucleoside 3’-phosphate [for review, see reference (21)1. Exhaustive hydrolysis of RNA results in formation of a resistant “core” composed of purine oligonucleotides terminated by a pyrimidine nucleoside 3‘phosphate. High concentrations of RNase slowly degrade poly A, but normally

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the enzyme does not release free purine nucleotides, nor does it hydrolyze purine nucleoside 2’: 3‘-cyeIic phosphates ( 2 2 ) . RNase hydrolyzes esters of pyrimidine nucleoside 3’-phosphates, e.g., the benzyl, methyl, ethyl, or 0-naphthyl esters (23, 24), but not the esters of pyrimidine 2’-phosphates or of purine nucleoside 3’-phosphates. Under suitable conditions, the enzyme exhibits synthetic activity, catalyzing the formation of alkyl esters from nucleoside 2‘: 3’-cyclic phosphates and primary alcohols, or of oligonucleotides from nucleoside cyclic phosphates and nucleosides (25, as), and this synthetic activity can be enhanced relative to the hydrolytic activity by suitable modification of the structure ( 2 7 ) . Goldstein (28) has shown that a specific alkylated derivative of RNase I, ~-carboxymethyllysine-41RNase, is devoid of synthetic activity and inert toward 2’:3’-CMP, but retains its endonucleolytic activity and specificity. A ribonuclease with a specificity similar t o that for the pancreatic enzyme has been isolated and purified 260-fold from KB cultured mammalian epithelial cells. However, the enzyme differs in two respects from that of the pancreatic type: its pH optimum is lower, p H 6 ; and it is somewhat less heat-stable ( 2 8 ~ )It. was not tested in high concentrations against poly A, the results of which would have been valuable for further characterization of the enzyme. AEkaZine RNases. Several ohservers (2953) have isolated from rat liver supernatant a relatively thcnnostahle enzyme, optimally active a t pH 7.5-8.0, that hydrolyzes RNA to resistant oligonucleotides with a terminal pyrimidine 2’: 3’-nurleotide, and that is inhibited by a substance found in the supernatant of liver and other tissues (19, 34, 3 5 ) . The properties and specificity of the inhibitor have been extensively reviewed (9). Roth (31) also reports the isolation from rat liver mitochondria of another alkaline RNase active toward RNA and pyrimidine nucleoside cyclic phosphates, but inert toward poly A, poly U, and adenosine 2’: 3’-phosphate, and inhibited by a supernatant inhibitor but not by the antiserum to crystalline RNase; the meaning of this is uncertain, since it has been shown (36) that the mitochondria1 and supernatant enzymes exhibit identical chromatographic behavior, pH optimum, and specificity. A chromatographically purified enzyme preparation from beef liver was found by Maver and Greco (37) to hydrolyze RNA to cyclic nucleotides, leaving a small core; exhaustive digestion led to the appearance of 3’-phosphates from the cyclic nucleotides. Beard and Razzell (38) purified alkaline RNase from pig liver mitochondria and supernatant 3000-fold and found the enzyme from both fractions to be identical. The enzyme hydrolyzes RNA more rapidly

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than synthetic polyribonucleotides, with poly A completely resistant, to liberate oligonucleotides with terminal 2’: 3’-phosphates. Activity against the cyclic phosphate ring could not be tested because of the presence of an inhibitor. The enzyme is not inhibited by antiserum to pancreatic RNase. RNase activity optimal a t alkaline p H has been found in many organs (39). Acid RNases (EC 2.7.7.17). Maver et al. (40) originally isolated from bovine spleen a heat- and acid-labile enzyme, with an optimum at pH 5.8, that degrades RNA to purine and pyrimidine nucleoside2’: 3’-phosphates. Purification showed the final products to be 3‘-phosphates (37), the 2’-phosphates previously reported being due to a very active contaminating 2 :3’-nucleotide phosphodiesterase (see below). Further purification gave an enzyme th at hydrolyzes tRNA and rRNA at almost equal rates, and poly U much more rapidly than other synthetic polynucleotides ( 4 1 ) . A similar enzyme ( S I - S S ) , subsequently shown to bc contaminated with a 2’: 3’-nucleotide phosphodiesterase (37), has been isolated from rat liver. However, Nodes et al. (36) used DEAE chromatography to isolate from rat liver mitochondria an RNase fraction inactive toward cyclic phosphates. This discovery was confirmed by Beard and Razzell (38) during the course of a study on acid RNase contamination of a hog liver alkaline RNase; after removal of a contaminating 2’:3’nucleotide phosphodiesterase, the acid RNasc exhibited little activity against nucleoside cyclic phosphates. Acid RNase-like enzymes are found in nearly all tissues, and are unaffected by natural (supernatant) inhibitors of alkaline RNases (9, 20, 51,SS).

RNases releasing 5‘-phosphate-temninated oligonucleotides (6’RNase) . Two such enzymes, both endonucleases, have been reported. One, purified from guinea pig (4.2) and pig (4.3) liver nuclei, hydrolyzes poly A to small oligonucleotides. It is optimally active a t pH 7, is activated by Mg2+, and hydrolyzes all polyribonucleotides except poly G (42). The other, with no marked base specificity, and isolated from the endoplasmic reticulum membranes of rat liver and Ehrlich tumor cells, is optimally active a t p H 7-8 and is insensitive to Mg” and N a F (44-46). 2. DEOXYRIBONUCLEODEPOLYMERASES DNase I (EC 3.1.4.5). This enzyme was first crystallized from the pancreas ( 4 7 ) , but similar activity occurs in other organs (16, 48-50), although no struetural identity between these has been established ( 9 ) .

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The enzyme is optimally active a t about p H 7 and is usually activated by Mg2+(49, 5 1 ) . Native DNA is depolymerized with a rapid decrease in viscosity; denatured DNA is attacked more slowly. The enzyme is inactive toward small oligonucleotides, esters of mononucleotides, apurinic acid, and deaminated single-stranded DNA ( 2 , 52, 5 3 ) . Many tissues contain natural inhibitors toward DNasc I (16, 54-57) ; two such protein inhibitors have been prepared in crystalline form from calf spleen (58, 5 9 ) . One of these, referred to as inhibitor 11, has now been identified as a labile protein with a molecular weight of approximately 60,000. It inhibits specifically DNase I and exhibits no adverse activity against DNase 11, snake venom phosphodiesterase, pancreatic RNase, or E . coli endonuclease ( 5 9 a ) . Under a variety of experimental conditions, in the p H range 6-9, it interacts directly with DNase I to form only one type of stable complex, with a ratio of inhibitor I1 to DNase I of 1:1 ( 5 9 b ) . Above pH 9.5 the complex dissociates with simultaneous denaturation of the inhibitor. The specificity, cation requirements, and nature of the products of DNase I action are still far from clear ( 8 ) , but studies by Bollum (60) with the aid of synthetic polydeoxyribonucleotides may lead to a resolution of these problems. DNase I I (EC 3.1.4.6). This enzyme, isolated from spleen (61) and thynius (62, 6 3 ) , occurs in nearly all mammalian cells (48, 50, 64, 6 5 ) , with an optimum a t p H 4.2-5.5 and maximum activity in rapidly multiplying tissues (66). The enzyme initially rapidly hydrolyzes about 10% of the internucleotide linkages, followed by a slow release of 3’-phosphate-terminated mono- and oligonucleotides ( 6 7 ) . Of considerable interest is the finding that the enzyme may split both strands in native DNA simultaneously to give two duplex strands, and the correlation of this behavior with its dimeric structure (68, 6 9 ) . DNase I1 is inhibited by Mg2+concentrations in excess of 1 mM and is activated by monovalent cations (49, 67); it is inhibited by denatured DNA ( 7 0 ) . It has been reported to exhibit some activity against bis-p-nitrophenyl phosphate and p-nitrophenyl esters of 3’-deoxynucleotides (71) . Tissue and urine inhibitors of the enzyme have been reportcd (2, 7 2 ) . DNase I1 from HeLa cells has now been purificd 700-fold ( 7 0 ) . Treatment of this enzyme with mercaptoethanol led to the production of two modified active enzyme species. Both of these, like the native enzyme, degraded native DNA by “single hit” kinetics. Kinetic data for both HeLa cell and calf spleen DNasc I1 suggested a “twin site’’ model capable of both “single hit” and “double hit” degradations.

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Lamb brain phosphodiesterase. This endonuclease preferentially hydrolyzes thermally denatured DNA. It has a broad p H optimum, 7-9, and leads to formation of 5’-phosphate-terminated oligonucleotides (73). 3. NONSPECIFIC ENDONUCLEASES

A thermolabile enzyme from chicken pancreas hydrolyzes DNA, RNA, and, a t somewhat slower rates, synthetic polynucleotides, with the release of small 5’-phosphate-terminated oligonucleotides. NO activity is manifested against nucleoside cyclic phosphates or esters of nucleoside phosphates. The enzyme is optimally active a t highly alkaline pH, is activated by divalent cations and inhibited by EDTA ( 7 4 ) . Rat liver mitochondria contain a nonspecific heat-labile endonuclease, optimally active a t pH 6.8; it hydrolyzes both RNA and DNA, with a marked preference toward single-stranded chains, to yield 5’-phosphateterminated oligonucleotides. It exhibits no base specificity and is inactive against p-nitrophenyl 3’- and 5’-TMP. The enzyme possesses an absolute requirement for Mg2+or Mn2+ (75, 7 6 ) . The relationship, if any, between this enzyme and another previously isolated from liver and purified fifteenfold in the same laboratory (77) is not clear. The latter endonuclease exhibits similar cation requirements and preference for denatured DNA. A similar heat labile endonuclease, with no base specificity or preference for the sugar moiety, optimal activity a t pH 7.0-7.5, and with a requirement for Mg2+,has been isolated from sheep kidney. The enzyme exhibits rigorous specificity with regard to secondary structure, attacking only nonstructured regions with formation of 5’-phosphate terminated oligonucleotides (77a). An additional enzyme hydrolyzing RNA a t alkaline pH occurs in the particulate fraction of rat liver ( 7 8 ) , but its specificity toward other substrates has not been examined. It is labile to heat and acid, with an optimum a t pH 9.CL9.5. It is not affected by rat liver supernatant inhibitor, is strongly inhibited by monovalent cations, and is activated by non-ionic detergents. 8. Exonucleases Phosphodiesterase I (EC 3.1.4.1). This enzyme, which resembles snake venom phosphodiesterase, is heat labile, optimally active a t p H 9.2, and is found in nearly all tissues (79-82). It is most active against chains with a 5’-phosphate terminal group, and it releases 5’-mononucleotides stepwise from the 3’-hydroxyl tertninal end. Acetylation of the terminal 3’-hydroxyl does not significantly affect enzyme activity, but chains terminating in a 3’-phosphate are relatively resistant. The enzyme is

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active against dinucleotides and nucleotide esters such as p-nitrophenyl or a-naphthyl thymidine S’-phosphate. It rather reluctantly hydrolyzes bis-p-nitrophenyl phosphate and dinucleoside monophosphates (83). Leukemic cell phosphodiesterase (EC 3.1.4.1). Isolated from mouse leukemic cells, this enzyme, optimally active a t p H 7-8, hydrolyzes oligo- and poly-, ribo-, and deoxyribonucleotides, with the stepwise release of 5’-mononucleotides. It is active against dinucleotides, but not nucleotide esters. It belongs to it newly discovered class of phosphodiesterases requiring the presence of two nucleoside moieties in ester linkage to a phosphate group ( 8 4 ) . Polynucleotide phosphorylase (EC 2.7.7.8). Although this enzyme is fairly widespread among bacteria, there are only several reports of its existence in mammalian cells (e.g., in rat liver nuclei and nucleoli) and exhibiting phosphorylytic activity ( 8 5 ) . Hilmoe and Heppel (86) isolated from guinea pig nuclei an enzyme converting poly A to ADP and catalyzing the exchange of P3‘0, with ADP; it is optimally active a t pH 7, requires Mg“, and is inhibited by fluoride. The discovery of such an enzyme in human sperm (87) has not been confirmed and is highly suspect. Phosphodiesterase IZ (EC 3.1.4.1). This enzyme is widely distributed in animal tissues (79, 88) and has been isolated from the spleen (89, 90). It is heat-labile, has an optimum a t pH 6, and releases 3’-mononucleotides from the 5’-hydroxyl end of the chain ; 5’-phosphate-terminated chains are resistant (90, 91). Phosphodiesterase I1 is active against some esters, like p-nitrophenyl thymidine 3’-phosphate, and may also catalyze formation of internucleotide linkages (26, 91). Mammary tumors of C3H mice contain an enzyme that produces 5’-mononucleotides from DNA, with a preference for denatured DNA (92, 9 3 ) . It is optimally active a t pH 8.5, with an absolute requirement for Mg2+,and is inhibited by Na+. Activity against p-nitrophenyl thymidine 5’-phosphate and RNA, to further define the specificity, was not reported.

C. Cyclic Nucleotide Phosphodiesterases 3’:5‘-iVucleotide phosphodiesterases. An enzyme found in various tissues, and most abundant in the brain, hydrolyzes 3’: 5’-ribonucleotides (3’: 5’-AMP) most rapidly, to 5’-phosphates, with optimum activity at pH 7.5-8.0 (94, 9 5 ) . It is activated by Mg?+ or Mn2+ ions, inhibited by methylxanthines (94, 95, 95a), nucleoside triphosphates, and inorganic pyrophosphate (95a, 95b), and is inert toward internucleotide linkages and nucleoside 2’:3’-cyclic phosphates ( 9 4 ) . Purified 170-fold from dog heart, it hydrolyzes 3‘: 5’-dAMP more rapidly than 3‘: 5’-AMP (96).

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Subsequently a second, analogous, enzyme was partially purified from dog heart. It hydrolyzes 3’:B-UMP a t a rate fourfold that of 3’:5’-AMP and 20 times that of 3’:5’-CMP, with a n optimum a t p H 8. By the use of ammonium sulfate, this enzyme was partially separated from the one described in the previous paragraph, and was found to be inhibited by 100-fold lower concentrations of methylxanthines. On the basis of this evidence, it is considered to be a different enzyme than that hydrolyzing 3’: 5’-AMP (97). It is quite conceivable that additional enzymes of this class may yct be found. 2’:s’-Nucleotide phosphodiesterase. Only one such enzyme has hitherto been reported. It is most abundant in nervous tissue ($7, 98-100), has been isolated from beef brain, and hydrolyzes purine and pyrimidine nucleoside 2’: 3’-cyclic phosphates to the 2’-phosphates, with a pH optimum of 6-7. It is inert toward internucleotide linkages, 3‘: 5‘-nucleotides, and nucleotide esters.

111. Methods of Assay The assay methods for nucleolytic c’nzymes outlined here are usudly applied in fractionation studies. However, they may and should be employed in determining the validity of histo- or cytochemical procedures, as is shown below.

A. Endonucleases I n the past, tissue RNases have been assayed almost exclusively against RNA substrates. However, the oligonucleotides initially released by RNases are subject to further degradation by tissue phosphodiesterases. A similar situation prevails for DNases. I n some instances, a suitable pH in the incubation medium may selectively inhibit phosphodiesterases. However, the usual assay of the acid-soluble products of what is believed to be RNase or DNase activity is a measure of both the endoand exonucleolytic activities of a tissue sample. It has been pointed out by Razzell (79) that the discovery of phosphodiesterases I and I1 in nearly all animal tissues renders dubious many of the earlier quantitative estimations of tissue RNase and DNase. 1. RNASES

Methods of measuring endonucleolytic RNase activity are generally unsuitable for tissue homogenates; this applies to the Kunitz (101) method and others, reviewed by Josefsson and Lagerstedt ( 5 ) and by Roth (9). The turbidimetric method has been claimed to be the only

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one generally applicable to crudc enzyme preparations (102). Assay of acid-soluble products formcrl in later stages of the reaction has been widely employed, although tliis is not always too accurate. The relative merits of various precipitating agents have been evaluated by Rot11 ( 9 ) . Furthermore, coniinercial RNA preparations, widely used in the past as the substrate, contain polyvalent cationic contaminants that may affect results by modifying enzyme activity directly (38) or by inactivating the inhibitors (34, 1031. Roth ( 9 ) circumvents this by addition of E D T A to the assay nicdium or by the use of purified, highly polymerized yeast RNA (104 ) , relatively insensitive to exonucleases. But it is important t o note that E D T A may unmask ribosome-bound enzyme activity ( 1 0 5 ) . Polyeytidylic acid, now commercially available, is a fairly sensitive substrate for RNases of the pancreatic type (106). Some increase in specificity may be attained by control of pH and ionic strength, or the use of activators and inhibitors, etc. Neu and Heppel (105) estimated the latcnt RNase I of Escherichia coli with tRNA in the presence of EDTA, i.e., under conditions mfavorable for E . coli phosphodiesterase and polynuclcotide phosphorylase. Free RNase was estimated using tRNA as the substrate in the presence of Mg“. Estimations may also be based on the relative rates of hydrolysis of various types of RNA or synthetic polynucleotides, and on the nature of the products formed (107, 108). It is rather difficult to obtain specific substrates for endonucleases. Pancreatic RNase, because of its biphasic mode of action, is an exception in view of its activity against pyrimidine nucleoside 2‘: 3’cyclic phosphates, but even it is difficult to assay specifically bccausc of susceptibility of the cyclic products t o 2‘: 3‘-nucleotide phosphodiesterases, which, as a result of their wide distribution and broad pH optima ( 9 9 ) , may interfere with RNase assays. Such interference can be detected by the formation of nucleoside 2’-phosphate products ( 3 8 ) . Hydrolysis of nucleoside cyclic phosphates may be followed by electrophoresis or by paper or thin-layer chromatography (5, 109, 110), but the rates of hydrolysis are low (Section VII). Titrimetric (5, 111) and spectrophotometric (5, 112) methods for following hydrolysis have been described but are difficult t o apply t o homogenates. Butcher and Sutherland (94) followed the hydrolysis of 3’: 5’-nucleoside phosphates by addition of a 5’-nurIeotidase, followed by colorimetric dctermination of the P, liberated. This procedure (cf. Scheme 31, originally suggested for RNase assay by Heppel [cf. ( l o g ) ] ,is applicable to any phosphodiesterase assay and niay be used with tissue extracts if the 5’-nucleotidase is replaced by a nonspecific phosphatase. Other useful substrates are nucleoside 3’-alkyl or aryl phosphates,

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although their susceptibility to phosphodiesterase I1 limits their application to alkaline RNases. After earlier attempts (11S), a diester, a-napthyl uridine 3’-phosphate was synthesized for this purpose. This suhstance yields a color-forming product on hydrolysis and, even more important, it is resistant to phosphodiesterase 11. The high rate of hydrolysis of this substrate jcf. Section VII, B) and its specificity make it very suitable for both colorimetric and histocheinical assays of RNase (114). Since alkaline RNases often occur in an inhibited form, the total, as contrasted with the free, activity is assayed in homogenates subjected to freezing and thawing in the presence of p-chloromercuribenzoate (17, 103). Acid RNases may be assayed with RNA “core” as substrate (115); the core is resistant to alkaline RNases, but not to phosphodiesterases. It is possible that a-naphthyl esters of purine nucleoside 3’-phosphates, resistant to phosphodiesterase 11, may be more suitable, and attempts are under way in our laboratory to prepare such a compound. RNases that lead to formation of 5’-phosphate-terminated products, like the K+-activated RNase or the liver nuclear enzymes, are generally assayed with poly A as substrate in the presence of the proper activating ions (42, 43, 46, 116). Free and latent acid RNase may be assayed together, in the presence of low concentrations of p-chloromercuribenzoate and with repeated freezing and thawing of the tissue extracts (10s). 2. DNASES

Methods of DNase activity assays and their applications have been reviewed by Kurnick ( 4 ) and Laskowski (8). The viscosimetric method, as well as that based on the UV hyperchromicity accompanying hydrolysis, both reflect initial cleavage by the enzyme, but are difficult to apply to homogenates ( 4 ) . A procedure that estimates exclusively endonucleolytic activity makes use of the loss of activity of transforming DNA ( 1 6 ) , or of A-phage infectivity ( 6 8 ) , but these are too complex and time-consuming for routine work. The methyl green method, which probably measures more than just endonuclease activity, is claimed to be applicable to alkaline and acid DNases of tissue homogenates ( 4 ) but has not gained many proponents. Assay of acid-soluble products, either by spectrophotometry or the diphenylamine reaction, has been most widely used in studies with crude systems. I n assaying DNase I, Mg2+ and pH control are used to discriminate against DNase 11, and conversely for DNase 11. A sensitive method for acid-

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soluble products, based on fluorimetry of thymine-containing oligonucleotides, has been proposed for acid and alkaline DNases ( 1 1 7 ) . Antiserum to DNase I has been employed for estimations of DNase activity in salivary glands of Chironomus thummi (lit?), but the specificity is questionable as the immune serum employed also precipitates DNase I1 and streptococcal DNase. Immunochemical assay of DNase preferentially attacking singlestranded DNA is highly specific, extremely sensitive, and free from exonuclease interference, since the serological activity depends on the size of the DNA molecule and hence responds to endonucleolytic breakage ( I 19). A rapid and sensitive method for endonucleolytic DNase activity (120), applicable to tissue extracts ( l d l ) , is based on the ability of nitrocellulose filters to retain only large fragments of denatured DNA. Labeled denatured DNA is incubated with the enzyme and then filtered; the decrease in retention of radioactivity is correlated with enzyme activity. For DNases with a preference for native DNA, the latter is denatured after incubation and then placed on the filter. Total DNase activity is assayed after dcstruction of inhibitors by aging or heat treatment (55, 56).

B. Natural Inhibitors Many tissue endonucleases are known to be associated with natural protein inhibitors, which may interfere with assay of enzyme activity. Methods have been elaborated for assay of the inhibited enzyme (see preceding sections) and of the inhibitor activity directly. Alkaline RNase inhibitor is estimated from the amount of tissue extract which gives 50% inhibition of crystalline RNase (9, 103), and similarly for DNase I (55, 57) and DNasc I1 (72).

C. Exonucleases Phosphodiesterase I . Earlier methods of assay of phospliodiesterase I were based on the use of bis-p-nitrophenyl phosphate, but this subbtratc is also susceptible to I)liospliodiesterase I1 and DNase I1 (71) and is hydrolyzed a t only 1% of the rate for the natural substrates, which are 5'-phosphate-terminated dinucleotides. Razzell and Khorana ( 8 3 ) introduced p-nitrophenyl thymidine 5'-phosphate, specific for 5'mononucleotide-forming exonucleases and hydrolyzed a t a rate six times that of the best natural substrates; the liberated p-nitrophenol is then easily estimated colorimetrically a t alkaline pH (6). Alternatively, a-naphthyl thymidine 5'-phosphate may be used but, apart from histo-

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chemical applications (122), it presents no special advantages over the p-nitrophcnyl analog. Finally, both a-naphthyl and 3-pyridyl thymidine 5’-phosphates undergo small spectral changes a t 3101 mp when liydrolyzcd ( I d s ) , but these changes are too small to be of practical usc except with purified enzymes and difference spectrophotometry. Phasphodiesterase II. Early assay methods made use of bis-p-nitrophenyl phosphate, but this proved a poor substrate; it is also subject to some interference from phosphodiesterase I (83) and, to a lesser extent, from DNase I1 ( 7 1 ) . One may use RNA “core,” with spectral estimation of acid-soluble products (115),but interference from acid RNases is to be expected. Razzell (79) introduced p-nitrophenyl thymidine 3’phosphate for enzyme assay of tissue extracts; it is hydrolyzed a t about the same rate as the natural substrates and the rate can be estimated colorimetrically ( 9 1 ) . Although very convenient, its trustworthiness is somewhat liinited by a slight susceptibility to splenic acid DNase ( 7 1 ) . Bernardi and Bernardi (41) consider 3’-phosphate-terminated oligodeoxyribonucleotides (resulting from exhaustive digestion of DNA by splenic acid DNase) more specific substrates for phosphodiesterase 11; these undoubtedly merit further study.

D. Cyclic Nucleotide Phosphodiesterases

2’: 3‘-Nucleotide phosphodiesterase. Hydrolysis of 2 :3’-AMP to 2’AMP is followed by measuring the Pi released from the latter by an excess of phosphomonoesterase (6). The use of purine nucleoside cyclic phosphates eliminates the interference of alkaline RNases. 3’:5’-Nucleotide phosphodiesterases. Specific substrates are the 3’ :5’cyclic nucleotides. Here again the method of choice is to follow the release of P, in the presence of added phosphomonoesterase ( 9 4 ) .

IV. Substrate Preparations I n view of the multitude of nucleolytic enzymes found in rnammalian cells (and there are even more in lower organisms) and the overlapping specificities frequently encountered among these, it is perhaps somewhat of an anachronism to speak of specific substrates, unless it is with reference to a given enzyme. However, even a substrate with limited specificity is still indispensable in any method designed to estimate or localize a given enzyme, or class of enzymes. The following paragraphs list some of the techniques for the preparation of substrates with absolute or relative specificities. Ribonucleoside 2’: 3’-cyclic phosphates have been synthesized by a variety of procedures, but we limit ourselves here only to those that do

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not require supplementary purification techniques. The method used in our laboratories for several years is based on the reaction of the commercially available nuclcoside 2’ and 3’-phospliatt~s with dicyclohexylcarbodiiinide (DCC) in anhydrous medium, to give the cyclic phosphates in quantitative yield in the form of the sodium or ammonium salts (124, 125). The method of Smith et al. (126‘) involves the reaction of the ammonium salts of the nucleotides with DCC in aqueous tertbutyl alcohol and formamide. Tha t of Michelson ( l 2 7 ) , based on treatment of the nucleotide in the presence of a base with ethylchloroforinate, has been made quantitative with the aid of a slight inodification by Taylor and Hall (128) for uridine 2’:Y-cyclic phosphate. Wigler (129) has described a procedure for obtaining cytidine 2’: 3‘-cyclic phosphate in crystalline form. Ribonucleoside 3’:5’-cyclic phosphates are conveniently prepared by reacting the ribonucleoside 5’-phosphoromorpholidate with DCC as described by Smith and Khorana (130). Additional procedures for all the natural ribo and deoxyribo analogs, including the widely eiiiployed 3’: 5’-AMP (now available commercially) have been described in detail (131). p-Xitrophenyl thymidine 3’-phosphate is synthesized by phospliorylation of 5’-O-tritylthymidine with p-nitrophenyl phosphorodichloridate (132).

a-Naphthyl phosphoryldichloride ,I

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p-ATitrophenyl thymidine 5’-phosphate may be prepared by esterification of thymidine 5’-phosphate with p-nitrophenol in the presence of

DCC (133).

The synthetic procedures for the p-nitrophenyl esters of all the ribo- and deoxyribo-nucleoside 5’-phosphates and 3’-TMP have recently been slightly improved upon and described in extensive detail (1S4). a-Naphthyl thymidine 5’-phosphate has been prepared by treatment of 3’-acctylthymidine with a-naphthylphosphoi-yldichloride according to Scheme 1 (122). The 3’ isomer may be prepared in an analogous manner froiii .Y-O-tritylthymidine. a-Naphthyl uridine 5’-phosphate is obtained by treatment of 2‘,5‘di-0-tetrahydropyranyluridine with a-naphthylphosphate in the presence of DCC (Scheme 2) (114).

4

OH ANN 3

OAN pyranylation

____c

d

O

I

ICSI

HOCH,

0

-80% CH,COOH

rG O-e>

OH

U-3’-p-Naphthyl

SCHEME 2

A modification of this procedure (114) makes possible the preparation of a mixture of the 2’ and 3’ isomers, which may then be separated from each other by several methods. Dinucleoside monophosphates, dinucleotides, and higher oligonucleotides. A variety of these is now available commercially, and the number

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of procedures for their chemical synthesis is too numerous to present here (see, e.g., 155-1373. Some of the ribo compounds can be very conveniently obtained on a preparative scale with the help of pancreatic or T 1 RNases by techniques described by Bernfield (138) and Egami (139).

V. Cellular Fractionation Endonuclease determinations have, with relatively few exceptions (see below), been based on assays of acid-soluble products released from RNA and DNA. Substrates with better specificity have been employed for exonucleases. Reference has already been made (Section 111) t o the question of substrate specificity and to the use of unpurified commercial substrates affecting cellular inhibitors. An additional problem, which has received little attention in fractionation studies, is the concomitant liberation of cellular inhibitors and their effects on the activity of various fractions. Control of enzyme diffusion has usually been limited by the use of suitable thermal and tonicity conditions to prevent disruption of cytoplasmic structures; sucrose solutions have been widely applied for this purpose, since they also limit aggregation of the particles. In some investigations, the validity of the findings has been cxamined by testing for adsorption of the isolated enzyme to the various cellular fractions, low adsorption being regarded as evidence for the reality of the activity of a given fraction. The initial, erroneous localization of DNA polymerase almost exclusively in the cytoplasm [cited in reference ( l 4 O ) l illustrates the types of errors arising from enzyme diffusion in aqueous fractionation studies ; subsequent nonaqueous fractionation revealed the high specific activity of the enzyme in nuclei (56,141) and its extractability with buffer explained the aforementioned erroneous results. Another example is that of LLribosomalRNase,” discussed in Section IX. The presently available wide variety of cell fractionation techniques and their applicability to intracellular enzyme distribution studies are discussed in several reviews (1&-145). The isolation of nuclei with the aid of nonaqueous media by Allfrey and Mirsky (1.46) represented probably the earliest attempt to avoid artifacts due to enzyme diffusion and concomitant rcadsorption in studies on localization of nuclease enzymes. Their procedure has been widely adopted and its grneral aspects were recently rrviewed (147). The use of nonaqueous media has also found application in the isolation of defined fragments from individual cells by microdissection, fol-

384

DAVID SHUGAR AND HALINA SIERAKOWSKA

lowed by incubation of the fragments with a nucleoside 2’:3‘-cyclic phosphate for RNase estimations, the extent of hydrolysis being determined by microelcctrophoresis ( 1 4 8 ) . Difficulties encountered in microdissection of small mammalian cells limit the procedure to relatively large invertebrate ooeytes, but improvements in microdisPection techniques could considerably broaden the potentialities of this technique. It is not without interest that microdissection of cells for such purposes was attempted as early as 1941 by Bundling (149)) who separated salivary gland nuclei from the cytoplasm in studies on catalasc distribution. Localization data from isolation procedures. In a number of instances, specific enzymes have been isolated from psrticular cellular fractions, e.g., the endonucleolytic RNase from guinea pig liver nuclei, which releases oligonucleotides with terminal 5’-phosphate groups (42). Since the emphasis in such studies has been on isolation and/or purification, and not on intracellular localization, these examples are cited in Section 11, and their possible significance is referred t o in Section IX. Brid RNases. Intracellular localization of nucleases by cell fractionation techniques was initiated in 1952 by Schneider and Hogeboom (150), who observed that the specific activities of acid RNA and DNA depolymerases of rat liver mitochondria markedly exceed those of the homogenate. Ultrasonic disruption of the isolated mitochondria liberates their depolymerase activity into the medium. The liberated enzymes exhibit little tendency to adsorption on mitochondrial structures, testifying to the authenticity of the mitochondrial localization. Subsequent investigations (33, 63, 151, 152) of acid RNase revealed that the specific activity of particulate fractions exceeds those of the nuclei and supernatant. I n liver the activity has been found mainly in the mitochondrial fraction (33,151).D e Duve e t al. (153-156) attempted further subfractionation of the particulate fraction by various methods, including sucrosc density gradient equilibration, and found the acid RNase to be located, together with other acid hydrolases, in a distinct class of cytoplasmic particles, the lysosomes. These were shown to be enclosed by a membrane that could be disrupted by osmotic shock, sonification, or non-ionic detergents, with concomitant liberation and activation of acid RNase (cf. Schneider and Hogeboom, above). Reid and Nodes (3%’)attempted the localization of enzymes of known specificity by comparing activities of particular fractions against RNA and nucleoside 2.’ :3’-phosphates, and distinguished two acid RNases in the liver. They concluded that the acid RNase of lysosomes, which hydrolyzes RNA and also adcnosinc 2’: 3’-phosphate to 2’-nucleotides, is probably different from the acid RNaee that hydrolyzes cyti-

LOCALIZATION OF NUCLEASES

385

dine 2’:3’-phosphate to the 3’ isomer, the localization of which was difficult to establish. This finding warrants reexamination, since liver acid RNase has now been shown to tw inert toward cyclic nucleotides (Section 11, A, 1 ) . Roth (157) found no acid RNase associated with rat liver microsomes or ribosomes. In the kidney, acid RNase has been located in droplets together with other acid hydrolases (158, 159). I n mouse pancreas, the enzyme was found mainly in the zymogen granule and microsoinal fractions (160, 161), the microsomal enzyme being subject to activation by EDTA (161). The kidney droplets and the zymogen granule fraction are considered to be examples of polymorphism of lysosomes (144). Spleen acid RNase was reported to be concentrated in the mitochondria, but this must be regarded with reserve since no separation of mitochondria from lysosomes was undertaken (15.2). Smith and Winkler (162) lysed the purified bovine adrenal gland chromaffin granule fraction by exposure to hypotonic media, thus releasing acid RNase and DNase activities. Subsequent sucrose density gradient distribution studies showed that the acid RNase and other acid hydrolases of bovine adrenal medulla occur in the lysosoinee (163). Using the fractionation procedure of Schneider and Hogehoom (150), Girij a and Sreenivasan (2020) localized acid RNase in the mitochondria1 fraction of rat adrenal gland and liver. Freezing and thawing or detergent treatment appreciably increased the activity without affecting that of alkaline RNase. Acid RNase of rat brain was recovered from mitochondria, microsomes, and supernatant, with a somewhat higher specific activity in the supernatant fraction. Density gradient subfractionation of the mitochondria and microsomes showed the enzyme to be concentrated in the most dense fractions (164). Acid RNase of liver nuclei has been reported to range from negligible (33, 35) to 1-5% of the total (150, 153, 154). It has also been found in ‘honaqueous” nuclei (165). In other tissues, such as kidney (33) or thymus and spleen (18, 6 3 ) , the nuclear content of enzyme is higher. Atkatine RNase has been localized in the mitochondria1 (20, 33, 35, 151) and microsomal fractions (3.2, 33, 35, 151), and in the supernatant fraction (20, 32, 33, 35, 151) of liver cells. Roth (9) suggests that the supernatant activity arises not from alkaline RNase, which is rendered inactive by excess inhibitor in the supernatant, but rather from other RNases unaffected by the inhibitor. Inactivation of the RNase inhibitor in the supernatant increased appreciably the percent activity in the latter fraction (20, 3 5 ) . Roth (157) found some alkaline RNase as-

386

DAVID SHUGAR AND HALINA SIERAKOWSKA

sociated with rat liver microsomes and ribosomes, but subsequent studies (166) showed the latter finding to be an artifact (Section IX). Belousova (167) has claimed predominantly mitochondrial localization of spleen alkaline RNase, whereas in the pancreas the enzyme was concentrated in the microsoIncs, zyniogen granules, and supernatant (160, 168). In rat adrenals the enzyme was present in the mitochondria as well as in the supernatant (20). Studies dealing with the occurrence and inactivation of alkaline RNase inhibitors (20, 34, 35, 103) have altered our notions as to the ratio of alkaline to acid RNases in most tissues. Destruction of RNase inhibitor, which is localized in the supernatant, increases the level of the alkaline enzyme severalfold. Since the mitochondrial alkaline RNase occurs in an inhibited form to a lesser degree, removal of the inhibitor modifies the intracellular distribution pattern of the alkaline enzyme. Roth (35) compared the activities of acid and alkaline RNases in various fractions of rat liver cells. By inactivating the alkaline enzyme inhibitor, he estimated the total alkaline activity of a given fraction. The content of acid RNase was then calculated from the fractional activity resistant to inactivation. The relation between activities of individual fractions and the assay pH suggests that, in contrast to other fractions, the nuclei contain small amounts of alkaline RNase and no acid enzyme (169). Siebert et aZ. (85) found alkaline RNase in rat liver nuclei and nucleoli, partially in latent form, the nucleolar enzyme exhibiting a higher specific activity than the nuclear. The alkaline RNase activity of “nonaqueous” nuclei of rat liver is appreciably higher than that of nuclei isolated in sucrose ( 1 6 5 ) . Alkaline RNase has also been found in “aqueous” nuclei of kidney cells (33). 5’-Ribonuclease (cf. Section 11, A, 1) has been recently assayed in rat liver fractions, with poly A as substrate (Table I ) . About 65% of total activity was associated with the mitochondria. Nearly 25% appeared to be localized in the nuclear fraction, but its low specific activity makes this finding suspect and due, possibly, to contamination from the mitochondrial fraction ( 1 0 8 ) . DNase I distribution studies on liver fractionated by differential (170) and density gradient (155, 171) centrifugation point to the occurrence of this enzyme in mitochondria, the specific activity of the nuclear fraction being much less (170) and that of the nucleolar lower still ( 8 5 ) . Predominantly mitochondrial localization was likewise found in mouse pancreas ( 1 7 2 ) . DNase I activity has also been studied in “nonaqueous” nuclei of rat regenerating liver ( 5 6 ) and of calf and rabbit thymus (173).Specific activity of such nuclei was lower than that of the “nonaqueous” cyto-

387

LOCALIZATION OF NUCLEASES

plasm and was largely resistant to extraction by buffer. Tlic DNasc I of regenerating rat liver nuclei was associated to a larger extent than the cytoplasmic activity with a powerful inhibitor, the removal of which resulted in a twelvefold increase in enzyme activity ( 5 6 ) .Calf and rabbit thymus nuclei also contain DNase I inhibitors (173). TABLE I INTRACELLULAR DISTRIBUTION OF PHOSPHODIESTERASE I A N D 5’-ltX.is~ IN

RAT LIVER,WITH

(iLlrCOSE

6-PHOSPHATASE

AND

PHOSPHATASE AS REFEREN(IE ENZYMES (108)

Enzymes

Numher of Ahsoexperi- lute merits valuesa

Proteins

4

5’-RNase

4

Phosphodiesterase I Glucose Bphosphatase Acid phosphatase

4

3 3

ACID

Percentage distribution*

W

nt

N

L

blc

S

Recovery

10.5 f2.1 3.7 f2.5 6.8 f6.2

17.0 k3.1 2.8 k0.8 35.9 f5.9

43.1 k2.8 5.7 k1.6 9.7 k6.7

101.7 k7.6 100.0 f22.9 99.9 k0.2

2.17 100 k0.72 3.01 100 k0.76

16 f 1 23 f 9 40 f 6

2

14 4 f0.7 64.2 k13.0 6.7 k2.6

20.49 100 k1.28

10.9 k6.4

9.0 +4.6

6.8 f4.0

69.9 flO.l

3.5 +1.6

100.1 +0.2

1.92 100

8.1 f2.5

14.8 k4.7

45.7 f10.5

20.3 k8.1

9.3 f1.8

98.2 k4.5

167

k 19

+0.27

100

7 7 G 1

x

Absolute values are in milligrams per gram of homogenate for proteins; in micromoles of AMP produced per rni1iut.e per gram of homogenate for 5‘-HNase using an extinction coefficient of 142 X lo8;i n mitw,moles of p-nitrophenyl5’-TMP hydrolyzed per minute per gram of homogenate; and in micromoles of phosphate liberated per minute per gram of homogenate for glurose 6-phosphatase and acid phosphatase. * To express the percentage distriliut,ion, the absolute values for the homogellate were taken as 100. H, homogenate; N, nuclei; M, mitochoiitlrin; L, lysosomes; Mc, microsomes; 6, superiiatant.

DiVase II was initially localized by Schneider and Hogeboom (150) and subsequently by others (6‘5, 174) predominantly in mitochondria of liver and other organs. Beaufay et al. (170) found the enzyme in the lysosomal fraction of liver cells, confirming this by sucrose density gradient centrifugation (155, l 7 1 ) , which has also been employed to localize acid DNase in the lysosomal fraction of bovine adrenal medulla (163). In the kidney it occurs in droplets, which are probably the biochemical equivalent of liver lysosornes (144) and are characterized by

388

DAVID SHUGAR AND HALINA SIERAKOWSKA

high specific activities of other acid hydrolases (158, 159). I n rat brain the enzyme is concentrated in mitochondria and microsomes, with the highest specific activity in the densest subfractions (164). Allfrey and Mirsky (146) localized DNase I1 in ‘Lnonaqueous” nuclei of various organs and found the ratio of nuclear to cytoplasmic activities to vary appreciably for different tissues, being highest with liver and heart. The higher values obtained, as compared to those from nuclei isolated in an aqueous medium, are probably due to enzyme diffusion from the latter into the aqueous medium. The enzyme content of nuclei isolated in an aqueous medium also varies with different organs (63, 65, 154, 170, 174). R a t liver nuclei isolated in the presence of Ca2+ ions (175) contained 14% of total cellular acid DNase, and this activity was released from nuclei on transfer to a calcium-free medium, but not by treatment that solubilized the lysosomes. The specific activity of rat liver nuclei was reported to exceed that of nucleoli (85). Phosphodiesterase I. An extremely useful study of the distribution of this enzyme in liver and kidney has been made by Razzell ( 7 9 ) , using p-nitrophenyl 5’-TMP as substrate. The enzyme was localized largely in the microsomes, with a relative specific activity manyfold in excess of that in other fractions (Table 11).Its occurrence in nuclei, as contrasted to adsorption to the nuclear membrane, could not be unequivocally established. De Lamirande et al. (108) reported 36% and 41% of total rat liver cell phosphodiestcrase I in the microsomal and nuclear fractions, respectively, with similar relative specific activities in both fractions, but negligible in others (Table I ) . Within microsomes, phosphodiesterase I was located in microsomal membranes but not in ribosomes ( 4 6 ) . The high nuclear activity is in sharp contrast with the results of Razzell (79) and those obtained by cytochemical methods (122). The enzyme has also been found in goat brain ribosomes (176). A phosphodiesterase active against bis-p-nitrophenyl phosphate a t pH 9 has been reported as an authentic component of the plasma membrane of liver cells. Its specific activity in the membranes was %fold that of the microsomes, the major contaminant of the membrane preparations. The membrane phosphodiesterase resisted extraction procedures that removed 75% of the membrane proteins ( 1 7 6 ~ )While . the activity a t pH 9 points to the foregoing enzyme as phosphodiesterase I, final classification must await tests against more specific substrates. Phosphodiesterase I I distribution in differentially fractionated liver and kidney cells of the rat has been studied with the aid of p-nitrophenyl thymidine 3’-phosphate, and was found predominantly in the supernatant (Table 11).In the case of hog kidney homogenates, density gradient centrifugation techniques pointed to the mitochondria1 localiza-

389

1,OCALIZATION OF NUCLEASES

tion of this enzyme, suggesting that the supernatant enzyme had been released preferentially from the mitochondria ( 7 9 ) . 3’:5‘-Nucleotide phosphodiesterases. With 3’: 5’-AMP as substrate, this enzyme has been assayed in tissues of the dog fractionated into a 2000 X g particulate fraction and supernatant, the latter being more active ( 9 7 ) . The corresponding particulate fraction from beef heart contained most of the activity, which was resistant to extraction in isotonic and hypotonic media ( 9 4 ) .Nair ( 9 6 ) found the total activity in dog heart to be equally divided between the 600 x g particulate fraction TABLE I1 INTRACELLULAR DISTRIBUTION OF PHOSPHODIESTERASE I A N D PHOSPHODIESTERASE I1 IN RAT LIVERA N D KIDNEY,USINGAS SUBSTRATES ~NITROPHEN 5’-TMP YL AND 3’-TMP, RESPECTIVELY (79) ~

~

~

~

Phosphodiesterase Sperific activitiesa Fraction

A. Liver fractions 1. Homogenate 2. Nuclei 3. Mitochondria 4. Microsomes 5. Supernatant solution B. Kidney fractions 1. Homogenate 2. Nuclei 3. Mitochondria 4. Microsomes 5. Supernatant a

b

Total unitsb

I

I1

I

I1

3.2 2.1 4.8 18.2 1.9

0.30 0.10 0.21 0.29 0.67

820 198 180 2 40 123

70.0 8.5 7.8 3.7 44.0

7.5 3.4 4.6 38.0 2.9

0.29 0.19 0.28 0.32 0.65

1120 91 123 670 150

59.0 4.1 8.8 7.2 34.0

In pmoles substrate/hr/mg prot.ein. In equal volumes of 10% homogeiiate in 0.25 M sucrose of each tissue.

and the 100,OOOx g supernatant; liberation of almost all activity from frozen tissue into the supernatant ( 9 6 ) probably excludes localization in nuclei. I n r a t brain the enzyme was found predominantly in mitochondria, followed by supernatant, microsomes, and nuclei in decreasing order of activity. Subfractionation of the mitochondria showed the enzyme in the cholinergic nerve endings; subjection of mitochondria to osmotic shock indicated association of the enzyme with the soluble synaptic neuroplasm ( 1 7 7 ) . Subsequent investigations (95a, I77a) , showed the major port,ion of the microsomal enzyme to be in the latent form, and to become un-

390

DAVID S H U G A R A N D HALINA SIERAKOWSKA

masked on addition of Triton X-100. The unmasked microsomal phosphodiesterase constituted more than one-half the total activity, the remainder being distributed almost equally between the synaptoplasm and the 100,000 x g supernatant ( 1 7 7 ~ ) . The 3': 5'-UMP phosphodiesterase of dog heart has been found largely associated with the 2,000 x g particulate fraction and, to a lesser extent, with the supernatant. The fractional activity in the 2000 X g supernatant from other tissues was higher (9’7).

VI. Histochemical Methods These are based on what has become known as the film-substrate technique, originally proposed by Daoust (178) for the localization of DNase, and subsequently RNase (179, 180). A tissue section is placed in contact with a film of gelatin containing RNA or DNA. After incubation, the film is stained with a basic dye, toluidine, to give reduced staining in those areas where enzymatic hydrolysis had occurred. I n essence, the method is an extension of one developed earlier for the histochemical localization of proteases with commercial photographic emulsions. The film-substrate method has since been successfully applied to other enzymes, e.g., hyaluronidase (181) and amylase (181, 182), which require niacromolecular substrates. The substrate film is prepared by dissolving the substrate in aqueous gelatin, spreading a few drops of this on a slide, and placing the slide on a level surface until gelation occurs. Thinner films may be obtained, with accompanying higher resolution, by allowing excess solution to drain from a vertically supported slide (179). The films are then fixed in formaldehyde to make them water-insoluble and resistant to proteases. I n the original procedure, an unfixed frozen section of the desired tissue is mounted on a glycerol-gelatin supporting pad on a second glass slide, and placed in contact with the substrate film. After incubation, the slides are separated, stained, and examined under the microscope. While useful for some purposes (see below), it is more advantageous to mount the freshly frozen sections directly on the substrate film, incubate in covered petri dishes saturated with water vapor, and flush the sections off with a stream of water; adjacent serial sections, fixed and stained, are used as controls (183, 184). Mayner and Ackerman (185) modified this further by coating the section on the film with glycerol-gelatin to prevent drying during incubation, but Daoust (184) found that this led to masking of some reactive sites. Since incubation techniques of 4-24 hours were required with this coating technique, it must clearly affect

LOCALIZATION O F N U C L E A S E S

391

enzyme activity adversely. Furthcrmore, some RNase sites identified by Mayner and Ackerman (185) have not been confirmed by others. The technique is obviously simple and readily reproduced, while the viscosity of the “incubation medium” helps to reduce enzyme diffusion. However, enzyme diffusion is of lesser significance here because of the low resolution which, despite several improvements, does not permit localization a t the intnacellular level. This could conceivably be improved by the use of thinner adhering films such as those employed in autoradiography ; and Daoust (184) has proposed that superior films might be attainable by means of Kohler’s centrifugal spreading technique. The specificity of the method is, of course, limited, since it is of necessity confined to the use of polymer substrates that will not diffuse out of the fixed film-substrate matrix (184). I n addition, since even short oligonucleotides are capable of binding basic dyes [see reference ( 1 3 5 ) , p. 4521, the resulting localization is usually that of a mixture of endo- and exonucleases. Some extension of specificity is feasible by the use of additional natural and synthetic oligo- and polynucleotides as substrates (183). The original method suffers from the disadvantage that it pcrniits of no pH control during incubation, nor of the use of activators or inhibitors. This difficulty may he partially circumvented by modification of the pH of the substrate film, or of thr glycerol-gelatin supporting pad, discussed in detail elsewhere (185).

FIG.1. RNA-gelatin film exposed t o a section of rat kidney. Activity in 1~os1111al convoluted tubules (ISO). X 30.

392

DAVID SHUGAR AND HALINA SIERAKOWSKA

FIG. 2. DNA-gelatin film exposed to a section of large inrestine of the rat. Activity in epithelial cells wit.h little activity in the lamina propria, submucosa, and the muscle layers (184). ~ 3 0 .

While substrate films are rendered resistant to proteases by formaldehyde fixation ( I & ) , it is nonetheless advisable to make use of substratefree films as controls, followed by staining with alkaline (pH 10) toluidine, which is a sensitive test for protease activity (186). Nonspecific changes in staining of substrate films occasionally require additional controls. Films made from alkaline gelatin solutions, free of substrate, stain with toluidine blue; on contact with a tissue section, the degree of staining decreases considerably with prolonged incubation (186). Some nucleases have been localized with substrate-film made alkaline to increase the solubility of the polynucleotide substrate (183) ; these obviously require appropriate controls. Ignorance of the foregoing has resulted in what is probably false positive localization of RNase (180) and other nucleases in the kidney outer medullary zone (183).

Representative examples of nuclease localization by the film-substrate technique are illustrated by Figs. 1 and 2. Additional data may be found in a review by Daoust (184).

VII. Cytochemical Procedures The general requirements for cytocheinical localization of enzymes have been reviewed in detail by several authorities in the field (187189). A few points of interest relating to nucleasc enzymes, reviewed in part elsewhere (190), are outlined below.

LOCALIZATION O F NUCLEASES

393

Substrates having the specificities discussed in Section 111 should be susceptible to the least possible number of enzymes. Specificity may occasionally be further limited by control of pH, ionic strength, and inhibitors or activators. “Substrate” specificity is not, of course, of importance in immunofluorescence techniques. Enzyme diffusion is less of a problem than in aqueous fractionation techniques because of the possibility of introducing fixation. Fixation may often effectively prevent enzyme diffusion, but it is rare indeed for a fixative to induce translocation of a n enzyme from one site to another. However, fixation may have a deleterious effect on a localized portion, or all, of the activity in a section; it is consequently desirable to examine staining intensity and localization with more than one fixation method. Absence of enzyme diffusion, absolutely essential for a fully satisfactory cytochemical technique, should be tested for by ( a ) prior incubation of sections, with no subsequent change in activity or localization on incubation with substrate medium, or (b) incubation of a highly active section in contact with one exhibiting low activity; if there is no diffusion, each section will exhibit normal activity and localization. It is equally essential to test for nonspecific adsorption of the final precipitate, particularly a t sites of enzyme activity. This may be done by adding to the full incubation medium an exogenous source of the enzyme under study, and should result in formation of a uniform precipitate over the section area. If, however, it leads to an intensification of the reaction at sonie given site, localization of endogenous enzyme a t this site, while not disproved, is at least suspect.

A. Irnrnu nofl uorescence Techniques As early as 1954 attempts were made by Marshall (191) to localize RNase and DNase by means of this technique, which is based on the antigenic properties of the enzymes, so that they will readily precipitate fluorescein-labeled antibodies. Tissue sections were fixed in a buffered solution of formaldehyde in aqueous dioxane to avoid enzyme diffusion from sections. Both enzymes were localized in the zyniogeii granules and apical cytoplasm of bovine pancreatic acini ; there was some diffuse staining of the cytqplasm of certain acinar cells, while thc nuclei and mitochondria were negative. The validity of Marshall’s results for RNase were subsequently questioned by Ehinger (192) on the grounds that no tests had been made of the homogeneity of the antibody preparations and that this was responsible for the diffuse staining in Marshall’s preparations.

394

DAVID SHUGAR AND HALINA SIERAKOWSKA

Full details of the immunofluorescence technique in cytochemistry are given in several reviews (193-196). The technique is undoubtedly capable of high specificity, but this may be both an advantage and a hindrance. Proper purification of the antibody will lead to localization only of its antigen. Furthermore, antigenically identical enzymes exhibit narrow ranges of organ and species specificities. The antigenic specificity of antiserum to rat pancreatic and liver RNases has been examined by Gordon (197), who found the anti-pancreatic RNase serum to inhibit the activity of pancreatic and spleen RNases, and only partially those of kidney and serum, but not that of the liver. The anti-liver RNase serum was inert against the pancreatic enzyme and marginally inhibited the spleen enzyme but was highly inhibitory against liver and kidney alkaline RNases. No inhibition of acid RNases was noted. This apparently high specificity circumscribes the applicability of the method to highly purified enzymes. It is to be expected that, with further developments in purification techniques for enzymes, the high specificity of the immunofluorescence technique will make i t a highly refined research tool. It should be capable of differentiating between nucleases of different molecular structures and, possibly, of varying functional significance and should be applicable, for instance, to following the transfer of an enzyme within the secretory cells. It cannot, on the other hand, portray the overall enzyme activity in a cell; but this is a small price to pay for the high specificity attainable, the more so in that this gap can be filled with the use of less refined techniques of more general applicability (114). Fluorescent anti-liver RNase serum was used by Gordon and Myers (198) to localize RNase in the liver and kidney of the rat. The enzyme was found in the cytoplasm of liver parenchymal cells, with nuclei negative, and in the cytoplasm of proximal convoluted tubules of the kidney. No activity was noted in other normal tissues, nor in four liver hepatomas and one cholangioma. Intracellular details were rather poorly defined; this could have arisen from the use of ethanol fixation, which is known to be accompanied, during incubation, by enzyme diffusion (190). The same technique was applied by Ehinger (192) to localize RNase in Carnoy-fixed frozen sections of rat and ox pancreas. Two types of localization were reported, but not both in the same section: ( a ) positive nucleoli and basal ergastoplasmic zone of acinar cells (Fig. 3) with nuclei and remaining cytoplasm negative; (b) positive apical zymogenic areas, together with nucleoli, and the remaining cytoplasm negative. Islets of Langerhans were negative. This study included rigorous tests

LOCALIZATION O F NUCLEASES

395

for specificity. Oddly enough, however, a prior incubation of sections in buffer solutions led to subsequent decreased staining with antibody. The author suggested that this may have arisen from enzyme diffusion, w11ic.h normaIIy is prevented by blocking and fixation of the enzyme by anti-

FIG 3 Immunofluoresccnce localization of RNase in rat pancreas Actikity in I.)asal ergastoplasmic zone of acinar ccllls and tlmr nucleoli, w ~ t hnuclei and reinamIng cytoplasm ncgative (192). ~ 2 2 5

body. However, this cannot bc tlie full explanation for, with rapid enzyme diffusion, fixation by antibody should have been equally rapid ; since this should have led to staining, I t IS curious that 3 hours of incubation were required. I n attempting to explain the predominantly basal ergastoplasmic localization, as contrasted with the zymogen granule localization principally found by previous obstwers, it was proposed by Ehinger (192) that this might be due to looser binding of the zymogen enzyme after Carnoy fixation. If so, this should have been shown by other methods. Such discrepancies between immunofluorescence and other cytochemical

396

DAVID SHUGAR AND HALINA SIERAKOWSKA

methods are not limited to nuclease enzymes; the localization of aamylase in the basal portion of acinar cells by immunofluorescence (199) is in disagreement with results obtained by other procedures (182). Pilocarpine stimulation was found to lead to a pronounced decrease in RNase content of rat pancreas, but only to a given plateau level beyond which higher doses were ineffective. The inference was drawn that there must be two types of binding of the enzyme, sedentary and “exportable.” This was correlated with the finding of a dual localization, shown by histoiminunofluorescence and changes of localization during the diurnal cycle ( 2 0 0 ) . Finally, attention should be drawn to Ehinger’s (192) observation that formalin fixation of pancreas abolishes the antigenic activity of the tissue enzyme. This was not so for the crystalline enzyme, the antigenic activity of which was only partially modified by formalin treatment, nor did i t appear to be the case with the formalin-based fixative employed by Marshall (191), referred to above. I n any event the discrepancies between these results and those of Marshall (191) and ZanKowalczewska et al. (114) will require further study.

B. Precipitate-Forming Techniques 1. ALKALINE RIBONUCLEASES

Probably the first attempt a t localization of nucleases by precipitateforming techniques was that of Zugury e t al. (201), based on the observation that lead salts of RNA and oligonucleotides are more soluble than the corresponding mononucleotide salts. Formol-saline fixed sections were incubated in a medium containing RNA and lead nitrate (for RNase) , or oligonucleotides and lead nitrate (for phosphodiesterase) , the resulting enzymatically liberated mononucleotides being precipitated as the lead salts. Appropriate controls were used to assess possible interference from endogenous phosphomonoesterases. Localization patterns were identical for both substrates, mainly in cells of the Islets of Langerhans and a narrow band in the apical region of the secretory acini. Apart from the lack of specificity of this method, it was subsequently shown (183) that mononucleotide lead salts are far from insoluble; a t the lead concentrations used, precipitate formation a t pH 8 requires a mononucleotide concentration of about 2 mM, and even more a t lower pH. Concurrently with the above, RNase localization was attempted with the use of what a t that time was considered a specific substrate, a pyrimidine nucleoside 2’: 3’-cyclic phosphate (190). The specificity of this has since been shown to be only relative by the discovery of 2’:3J-

397

LOCALIZATION O F NUCLEASES

nucleotide phosphodiesterase (Section 11, C) . The scheme devised is essentially a two-step enzymatic reaction, the second step of which is based on the Gomori techniquc for alkaline phosphatase, as shown in Scheme 3. 4

*P:O

HO

PY

tissue RNase

i I

OH

HO

OH

0

0

exogenous alkaline phosphatase

i

HO-P-OH II 0 Po:-

Calcium phosphate

SCHEME 3

The incubation medium coiitained 2’:3’-UMP, purified alkaline phosphatase and a soluble calcium salt. Hydrolysis of the cyclic phosphate a t RNasc sites liberated uridine 3’-phosphate, which was dephosphorylated by the exogenous phosphatase ; the liberated phosphate ions were trapped by calcium to form a calcium phosphate precipitate, which was then revealed by the usual Gomori procedure. A rather serious disadvantage of the foregoing is the slow rate of hydrolysis of cyclic phosphates by RNases [Section I1 and reference ( 2 0 6 ) ] ,so that unduly long incubation times are required. Equally disadvantageous was the use of ethanol- and acetone-fixed sections, from which RNase diffuses rapidly [cf. reference (203)1, leading to artifactual nuclear and nucleolar localization. Enzyme diffusion probably could have been reduced to permissible limits by the use of formalin fixation (see below), but this was not attempted a t the time because of the claim (204, 205) that RNase in pancreatic sections was totally inactivated by formalin. Further work on this method was subsequently abandoned when nucleoside 3/-naphthyl phosphates were found to be more suitable substrates (see below) , but the above procedure undoubtedly warrants reexamination with the use of formalin and other methods of fixation, and reference should be made to the original paper (190) for a dctailed discussion of the problems involved in nuclease localization.

398

DAVID SHUGAR AND HALINA SIERAKOWSKA

A technique similar to the above utilizes RNA as substrate (206). Blood smears fixed in formalin vapor were incubated in a medium containing RNA, acid phosphatase, and lead nitrate. Activity was observed in all neutrophilic cells, mainly in the perinuclear region, and in lymphocytic cells, while controls without substrate and with addition of sodium fluoride were negative. However, no suitable controls were run for endogenous phosphatase, or for tissue affinity for the final lead phosphate precipitate. This procedure was subsequently applied to a study of the changes in RNase activity in leukocytes of rabies-susceptible and refractory mice; rather surprisingly, no RNase could be detected in the latter (207). Much more promising, and widespread in scope, was the development of the nucleoside naphthyl phosphates: a-naphthyl uridine 3‘phosphate (see Section I V and ref. 114) is completely hydrolyzed by RNase t o 2’:3’-UMP and free naphthol (see Scheme 4) ; its relative specificity for RNase is enhanced by the resistance of a-naphthyl nucleoside 3’-phosphates to spleen phosphodiesterase (122), which hydrolyzes other esters of nucleoside 3’-phosphates (91). The liberation of 0

y$ 0

HOCH,

H

0 OH I O=P-OH I

RNase

+

-

4 o, Ofl.-OH

+Naphthol

0

Insoluble red dye

5-Chloro-atoluidine

SCHEME 4

LOCALIZATION O F NUCLEASES

399

free naphthol accompanying hydrolysis by RNase logically suggested a one-step procedure for RNaee, based on coupling of the enzymatically liberated naphthol with a suitatilc djazotatc according to standard ~ Z O dye coupling techniques, as sliown i n Scheme 4. The utility of this substrate was further indicated by the fact that it is hydrolyzed by RNase a t w rate about 100-fold greater than that for 2’: 3’-UMP. The rapid rate of hydrolysis of a-naphthyl uridine 3’phosphate was further confirmed in collaboration with Witzel (unpublished rcsults), who found that I-,,,,, for this substrate was 80 times that for the corresponding benzyl ester, with the K , for the naphthyl derivative higher than that for 2’: 3’-UMP and uridine 3’-benzyl phosphate. Such a high rate of hydrolysis of the naphthyl derivative is probably caused by the high leaving tendency of the naphthyl group, which is reflected by the slow spontaneous hydrolysis of uridine 3’-naphthyl phosphate in aqueous medium. This slow rate of decomposition, leading to formation of free naphthol, would normally prove rather awkward in cytochemical work, but it is more than offset in this case by the rapid rate of enzymatic hydrolysis, making possible very short incubation periods. The slight, diffuse, nonspecific precipitate from spontaneous hydrolysis is readily removed during the normal rinsing procedure that follows incubation. The availability of this substrate led to a reexamination of the problem of enzyme diffusion, with the finding that cold formol-calcium fixation yields tissue sections with high RNase activity and m signs of enzyme diffusion (114). Previous reports (203-205) on the deleterious effects of formalin fixation a t 24” were confirmed, but fixation in Baker’s fonnol-calcium a t 4” led to little loss in activity, with either uridine 3’-naphthylphosphate or 2’: 3’-UMP as substrates. Activity against RNA was affected to a variable degree, depending on the effectiveness of foriiialin removal prior to incubation ; decrease in activity against RNA apparently resulted from interaction of residual formalin with RNA, rendering the latter more resistant to RNase. Cytochemical reactions were conducted at pH 9 to ensure a rapid coupling reaction and minimum diffusion of the reaction product. Frozen sections fixed in formol-calcium were routinely employed, with incubation times of 2-30 minutes. Extensive tests, including assays of incubation media for enzyme activity, established the absence of enzyme diffusion. The type of localization obtained is illustrated in Figs. 4 and 5. It is perhaps worth mentioning that with the incubation times employed no nuclear or nucleolar localization was ever observed (114). By contrast, and in agreement with previous observations (190), preparations of tissues fixed in acetone, alcohol, or Carnoy reagent ex-

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DAVID S H U G A R A N D H A L I N A SIERAROWSKA

hibited rapid and appreciable diffusion of enzyme into the incubation medium, which could be revealed by a variety of techniques. Such sections exhibited desmoactivity, which persisted following preincubation in water, and varied in different tissues from nil in the pancreas to a sizable percentage in the kidney. I n all cases investigated, activity of formalin-fixed material was far greater than the desmoactivity in alcohol or acetone fixed material, with the sole exception of the

FIG.4. Alkaline RNaae in rat pancreas by azo-dyc coupling technique. Intense activity in the suprannclear portion of acinar cells with lower activity in the remaining cytoplasmic area. Intense activity in the lumen of excrrtory ducts and slight activity in the epithelium of the excretory ducts adjoining the lumen (114). x 375.

vascular connective tissue enzyme. It has not been established whether the tissue-bound desmoactivity (located, for instance, in the kidney tubules) comes from pancreatic RNase differently bound to some tissue component(s) or, perhaps more likely, is due to a different RNase. It should be recalled that RNases differing immunologically from the pancreatic have indeed been detected in several organs, including the kidney (197).It would be desirable to determine whether the ratio of pancreatic to other alkaline RNases in a given organ can be correlated with the ratio of lyo- to desrnocomponent following acetone fixation. Should this prove

LOCALIZATION O F NUCLEASES

40 1

FIG.5. Alkaline RKase in rat kidney by aso-dye coupling technique. Intense activity in the brush border zone of thr proximal convoluted tubules (11 4 ) . X 1OOO.

to be so, it would imply a different functional significance for the two enzymes. This problem could Irofitably be tackled with the aid of iiiiniunohistochemical techniques. 2. DEOXTRIBONUCLEASES

Localization of DNase was first reported by Marshall (191), who used the histoinirnunofluorescence technique t o localize DNase I in the pancreas. The enzyme was found to occur mainly in the apical portion of acinar cell cytoplasm. No further work along these lines has since been done, possibly because of the antigenic nonhomogeneity of the enzyme. Precipitate-forming techniques have formed the basis of other methods, hit these h a w been somewhat limited in scope because of the absence of known simple substrates for these enzymes (see Section 11, A, 2 ) . A method for DNase I1 has been described (208) using D N A as substrate, with exogenous phosphatase in the incubation medium to liberate terminal phosphate groups exposed by tissue DNase. The libcrated phosphate was trappecl with lead ions and visualized by the Gomori technique. Substrate specificity is similar to that in other biochemical methods for DNasc I1 with DNA as substrate and elimination of othcr activities by pH control and a magnesium-free medium.

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DAVID SHUGAR AND HALINA SIEBAKOWSKA

DNase I1 activity, in frozen sections fixed in acetone-formalin-water and incubated for several hours, was reported in the nuclei and cytoplasm of splenic red pulp, in Kupffer cells, in nuclei of liver parenchymal cells, and in nuclei of the epithelium of kidney tubules and duodenum. These results must, however, be accepted with caution because of the following considerations. DNase localization was observed only in cells containing acid phosphatase. Furthermore, when exogenous phosphatase was omitted from the incubation medium, the same localization, but with decreased intensity, was observed ; while the exogenous conimercial phosphatase employed almost certainly contained phosphodiesterase activity. The long incubation periods required may have been accompanied by enzyme diffusion into the incubation medium [ cf. reference (190) 1. Finally, the nuclear localization observed was likely due to affinity of the nuclei for the lead phosphate precipitate released by the diffusing enzyme [cf. reference (189), p. 433, and reference (209)1. I n a subsequent application of this technique to tissues of irradiated animals (210) the fixation procedure was altered to include neutral formol-saline fixation of tissue blocks, washing in acetate buffer, dehydration with acetone, and paraffin embedding. This procedure was again modified by Vorbrodt ( a l l ) , using cold formol-calcium fixation, and a lower concentration of DNA and Pb2+in the incubation medium. The resulting localization was found to be pH dependent. It was mainly nuclear a t about pH 5, and largely in cytoplasmic granules a t about pH 6. The activity of cytoplasmic granules was inhibited by sulfate ions, which are known to inhibit DNase 11. It was also noted by Vorbrodt (211) that substitution of a commercial low molecular weight DNA preparation for the high molecular weight substrate resulted in an increase in activity. This raises some doubts as to the nature of the enzyme localized. Highly polymerized, native, DNA is subject to hydrolysis primarily by endonucleolytic DNases, but a degraded preparation is more susceptible to exonucleases. The Vorbrodt modification was subsequently applied by Coimbra and Tavares (212) to localization of DNase 11, as well as acid RNase (with an RNA substrate) in rabbit nerve tissue, with essentially similar pH-dependent results. Samorajski et al. (213) applied the foregoing to light and electron microscope localization of DNase I1 in dorsal root ganglia and the spinal cord of aged aniiiials, using 4-hOur incubation pcriods. The presence of DNase 11, acid phosphatase, and eathepsin-type-C estcrase reaction products in lysosomes and lipofuscin pigment masses was regarded as indicative of the role of lysosomes in formation of lipofuscin pigments.

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Nuclear DNase I1 activity was also reported in neurons. It is clear that this entire procedure requires critical reexamination. 3. PHOSPHODIESTERASES Phosphodiesterase I . Apart from the trials carried out by Zugury et al. (20f),referred to above, cytochemical procedures for phosphodiesterase have been proposed and applied with success (82, 122). One enzyme localized was phosphodicstcrase I, using as substrate the synthetic a-naphthyl thymidine 5’-phospliate (Section IV) . The enzymatically liberated naphthol is couplcd with a diazotate to form an insoluble colored precipitate, as described above for RNase (see Scheme 5 ) . The substrate is relatively specific for phosphodicsterase I and is hydrolyzed by this enzyme a t tlic same rate as the corresponding p-nitrophenyl ester (2f4), i.e., about 5 tiriics as rapidly as the natural substrate, a 5’-phosphoryldinuclcotide. This high rate of hydrolysis made possible relatively short incubation periods. Cold formol-calcium fixation of tissue sections reduced enzyme activity t o about 20%, with no subsequent diffusion, as estimated from both cytochemical and paper chromatography tmts. Paraffinembedded, acetone- or ethanol-fixed sections retained only 5% of the original activity, but with unchanged localization patterns. Enzyme activity was optimal a t about pH 9, but some activity persisted even at pH 5.2. Phosphodiesterase I activity obtained with formol-fixed sections is illustrated in Figs. 6 and 7. A consistent lack of nuclear and nucleolar activity is apparent in all the tissues examined. Attempts to apply the above procedure to localization of phosphodiesterase I1 with a-naphthyl thymidine 3’-phosphate (Section IV) , an analog of p-nitrophenyl thymidine 3’-phosphate ( 8 3 ) , were abandoned when the former was found to be totally resistant t o this enzyme ( 1 2 2 ) . 3’:5‘-Sucleotide phosphodiesterase. In an extension of the method described above for the localization of RNase by means of ribonucleoside 2’: 3’-cyclic phosphates, Shanta et al. (215) developed a two-step reaction for localization of 3’: 5’-nucleotide phosphodiesterase. In addition to the substrate, the incubation medium contained snake venom as a source of 5’-nucleotidase, and Pb2+ions to precipitate the liberated phosphate by the Gomori procedure as shown in Scheme 6. Appropriate controls appear to satisfy the normally accepted criteria for cytochemical localization (Figs. 8 and 9 ) . Omission of snake venom (i.e., 5’-nucleotidase) was accompanied by some activity, in rabbit liver, obviously from endogenous phosphatase, but with different localization, mainly in nuclei arid nucleoli. I n the presence of snake venom, no

404

d

i-

bZ

0

I

o=Pl-0

I x I

0

I

DAVID SHUGAR AND HALINA SIEXAKOWSK.4

sA38

\

V d

I

i x o=b-o

405

LOCALIZATION OF NUCLEASES

FIG.6. Phosphodiesterase I in rat liver by azo-dye coupling techniqae. Intensc activity in proximity of bile canaliculi. Cytoplasm of hepatic cells weakly artivc with somewhat grcater activity a t thc lumen of sinusoids; activity in capillaries ( 1 2 2 ) . x300

nucleotidePDase

OH

O=P-O I HO

+.PI

0

~ 0 -IIp - 0 - c ~ ~0

tissue

I

OH

%$ HO

p0,S-

+

5’-nucleotidase exogenous L-

lead acetate

+

Lead phosphate

SCHEME 6

HOC@*

HO

OH

OH

406

DAVID SHUGAR AND HALINA SIERAKOWSKA

FIG.7. Phosphodiesterase I in rat tongue hy azo-dye coupling technique. Iiitenec activity in t,he cytoplasm of mast rells, in the intima of small blood vesels, and in capillaries, with less activity in the perineurium (122). x 100.

FIG.8. htlcnosinc 3': 5'-~yelic phosphatc d k t e r a s e in rabbit liver by technique of Shanta ef nl. (215). Moderately positive activity in the cytoplasm of parenchymal cells with nuclei and nucleoli negative. x260.

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FIG.9. Control to Fig. 8 incubatcti in medium containing no snake venom. Thc localization corresponds to endogenous p1ioPy)honionoesterase and sites of high affinity for lead phosphate precipitiite. Not? the darkly stained nuclei and nucleoli (arrows) ~ 4 1 6 .

nuclear or nucleolar localization was observed, suggesting (a) that in the absence of snake venom, dcl)liosl,liorylation of AMP came from endogenous phosphatase, which diffused into the incubation medium, as occasionally happens, and that tlic rcsulting lead phosphate, liberated in the incubation medium, was adsorbed by the nuclei and nucleoli in accordance with their known affinity for these structurcs (189, 209) ; and (b) that in thc presence of snake venom, the resulting 5’-nucleotidase activity was high enough to override the effect of diffused endogenous phosphatase. Supplementary tests indicated no diffusion of cyclic nucleotidc phosphodicstcrase from the unfixed tissue sections employed in this study. Snake venom is known to hydrolyze slowly nucleoside 3’ :5’-cyclic phosphates; e.g., 3’: 5’-AMP is hydrolyzed by high concentrations of Crotalus adamanteus venom to give about 50% 3’-ARIIP and 50% adenosine, the latter resulting from dephosphorylation of 5’-AMP by the venom 5’-nucleotidase (216). This apparently did not interfere in the investigation discussed above since inactivated control sections were completely blank. Nonetheless, the replacement of snake venoiii by purified 5’-nucleotidase or highly purified nonspecific monophosphoesterase would be a desirable improvement.

C. Comparison of Cytochemical and Fractionation Findings A comparison of cytocheniical results with those of cell fractionation techniques is not a simple matter. Cytochemical procedures at present

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DAVID SHUGAR AND HALIN.4 SIERAKOWSKA

differentiate bctween specific regions within the cytoplasm or the nucleus, with the resolution of thc light microscope. Cell fractionation techniques locate enzymes a t an intraeellular level, i.e., in mitochondria, microsomes, lysosomes, ribosomes, more recently membranes, etc. With the current use of density gradient centrifugation and the perspectives opened up by zonal centrifugation ( 2 1 7 ) , the resolution begins to approach that of the electron microscope. The present availability of several suitable cytocheniical substrates emphasizes the need for application of electron microscopy to the cytochemical localization of nucleolytic enzymes. Efforts are already being made in the preparation and screening of coupling agents yielding electron-dense precipitates, by incorporation into diazotates of heavy metals (218) or osmiophilic radicals that, upon reaction with osmium, become electron dense (219). Either of these would be useful with the existing naphthyl derivatives. There is also the possibility of introducing electron-dense moieties into the precipitable component of a substrate. Alternatively, labeled naphthyl, combined with autoradiography, might serve for quantitative electron microscope localization. However, until these possibilities become realities, there are several lines of approach for improvement of existing methods. One of these is the search for additional specific substrates, as well as increasing the specificities of existing substrates with the aid of natural and synthetic inhibitors. Localization of natural inhibitors, which has hitherto received little attention, should be of value in elucidating their physiological functions (and that of the enzyme that each specifically inhibits). It should be perfectly feasible to localize such inhibitors by immunofluorescence techniques or, perhaps more simply, by fluorescein labeling of the appropriate enzyme. Quantitative evaluation of enzymatic activity is feasible with the use of appropriately labeled substrates or trapping reagents, in conjunction with autoradiography ( 2 2 0 ) . There remains also considerable room for improvements in trapping reagents, along the lines so extensively exploited, e.g., by Holt (221) for localization of esterases with the use of variously substituted indoxyl radicals.

VIII. Nucleolytic Enzymes in Pathological States Possible changes in levels of nucleolytic activities in tissues and body fluids in a variety of pathological states have been intensively investigated. There are, however, few, if any, instances where the results are sufficiently clear-cut t o permit definite conclusions. Some typical findings have been summarized ( 4 , 2 2 2 ) .

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The major difficulty in interpretetion stems from the fact that the variations encountered usually lie within the range of activities exhibited by the subjects in their noriuztl physiological state. It is possible that closer attention to the problem of cnzyiiie specificity in such studies may prove more profitable. Similar extensive investigations have been devoted to the influence of partial or whole-body X-irradiation on nuclease activities in diverse organs, occasionally in conjunction with estimations of other enzymes. The overall findings are too complex to sumniarize here and reference should be made to several reviews (223-225). The influence of drugs on nuclease activities has likewise been examined. For example, corticostcroids are known to provoke involution of lymphoid tissue, and this is associated with an increase in nuclease activities. Wiernick and MacLeod (266) noted significant increases in thymus DNase I1 and alkaline and acid RNases following injection of 9-a-fluoroprednisolone. The effect of the steroid on alkaline RNase disappeared aftcr removal of alkaline RNase inhibitor, implying that the initial action is on the inhibitor. This is, however, to be contrasted with the observation that the increase of adrenal gland RNA resulting from ACTH treatment can be attributed at least in part to elevated levels of RNase inhibitor ( 6 2 7 ) . We return to this subject below. It is worth adding, in this context, that streptococcal DNase (streptodornase) and, to a lesser extent, pancreatic DNase I (because of its commercial availability) have themselves found applications in clinical practice, vie. via local infusion or intravenous injection for the purpose of liquefying insoluble DNA-protein in purulent exudates in local infections. These applications have bcen reviewed in detail by Tillett (628). On theoretical grounds, oiie might rlxpect that a study of nuclease activities following virus infection would contribute more to an understanding of the possible regulatory role of these enzymes. The finding that, as with bacteriophages, virus infection is associated with the appearance of induced polymerascs has recently led to a search for other induced enzymes involved in the metabolism of nucleic acids. An alkaline DNase active against denatured DNA in HeLa cells has been reported to double in activity following vaccinia virus infection (629), the induced character of the increased activity being inferred from the arrest of the latter on addition of puromycin or infection with ultravioletirradiated virus. Induced DNase activity has also been claimed in cultures of baby hamster kidney cells infected with herpes simplex virus (250) ; the criteria for the induced character of the enzyme activity were based on heat stability, behavior toward native and denatured DNA, and response t o cations. Somewhat more convincing than the above is the

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DAVID SHUGAR AND HALINA SIERAKOWSKA

demonstration that pox virus infection of HeLa cells is accompanied by the appearance of several new enzymes active against DNA (231, 239). Since thcse were not detected in the prcsence of puromycin or fluorodeoxyuridine, their induced character, as coritrastcd to activation of latent activity, was presumed. The induced enzymes, two of which are exonucleases, differ from the corresponding enzymes in the normal cell either in type of attack or preference for native or denatured DNA. It is clear that further evidence for the induced nature of these enzymes, which should be directed more to such factors as specificity and type of attack, is highly desirable from the point of view of establishing their role in the infective process. Nucleases in Tumors This is a field that has attracted widespread attention. The results of earlier studies have been summarized in several reviews (9, 233, 234), from which it appears that there is a general tendency toward a lower activity of these enzymes, as compared to those in normal cells, for such tumors as ascites, hepatoma, and carcinoma of human cervix. However these findings are by no means uniform, one of the difficulties in comparing results of different observers being the lack of any uniform baseline for quantitative measurements of activity. Histochemical (film-substrate) methods have been applied in this field by Daoust et al. (235,256), who showed that azo-dye carcinogenesis in the rat leads to a progressive decrease in liver RNase before the parenchymal cells become cancerous; by contrast the loss in DNase activity is abrupt and appears to be closely associated with the neoplastic transformation of parenchymal cells (Figs. 10 and 11).A further study on 32 different types of 65 experimental and human tumors corroborated the earlier results for neoplastic cells, whereas the connective tissue stroma and necrotic regions exhibited high levels of activity (237), in accord with the generally observed increase in nuclease activity during tissue regression and necrosis. The foregoing results are somewhat a t variance with those of Roth e t al. (238) on a series of transplantable rat hepatomas and normal rat liver, using fractionation techniques. The original paper should be consulted for full details, but it is worth noting here that the hepatomas exhibited a doubling of the microsomal alkaline RNase activity and an appreciable increase in supernatant acid RNase. The discrepancy betweeen the results of Daoust et al. (235-237) and Roth et al. (238) may be more apparent than real, in view of the difference in techniques. Daoust e t al., using the film-substrate method, with an unbuffered gelatin supporting pad for the sections, had no control

LOCALIZATION OF NUCLEASES

41 1

over the pH, and hence detected probably a mixture of different RNases. Thc state of the enzyme in a tumor cell might also be such that its activity in a section is masked. It is indeed curious that this should be so for both RNase and DNase, but the findings of Daoust et al. (635-237) cannot be readily dismissed.

FIG.10. RNA-gelatin film exposed to a section of cirrhotic liver. Note t,he high ItNase activity in trabeculae of bile ducts and connective tissue. The nodules of pwenchymal cells are relatively i n d i v e (184). X 30. FIG. 11. R N A - g e l a h film eqiosed to a section of primary hepatoma. Note RNase activity in bands of connective tissue and the relatively inactive neoplastic cclls (T)(18.4). x30.

Phosphodiesterase I localization in unfixed frozen sections of human breast carcinomas and fibroadenomas has been studied by Michalowski and Jasinska (unpublished results), using a-naphthyl 5’-TMP. No activity was observed in the cancer cells, apart from occasional traces of activity in the necrotic regions. There was, on the other hand, intense activity in bands of connective tissue immediately surrounding, as if cncapsulating, the nodules of neoplastic tissue. The enzyme appeared t o be localized extracellularly, but did not show up uniformly in all connective tissue adjacent to caiicer nodules. Connective tissue bands further removed from neoplastic nodules exhibited little activity (Fig. 12) . The epithelial cells in fibroadenomas behaved erratically, some being negative, other exhibiting very intense activity in some of the cells. Vorbrodt (239) examined the cytochcmical localization of DNase I1 in various animal and human tumors and found the lysosomal activity in neoplastic cells diminished by comparison with that in normal cells.

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DAVID SHUGAR AND HALINA SIERAKOWSKA

Daoust (236) reported low activity against DNA films in neoplastic nodules with high activity in necrotic regions. Linked to the foregoing was the observation of Roth et al. (238) of a decrease in alkaline RNase inhibitor in rat hepatomas. More important still is the finding that mouse ascites tumor cells contain a potent RNase inhibitor most active a t neutral pH, leading to apparent pH optima of enzyme activity at acid and alkaline p H values, where it dissociates

FIQ.12. Phosphodiesterase I in human breast carcinoma by azo-dye coupling technique. Note the ncgativc cancer cells and the intense activity in bands of connective tissue adjacent to cancer nodules. x 100. (A. Michalowski and J. Jasinska, unpublished.)

from the enzyme (240). Of no less significance was the demonstration by Gordon (197) that RNases extracted from 5 tumors derived from liver are not inhibited by the anti-liver RNase serum. Since fluorescent antibodies t o rat liver RNase that stain normal liver intensely did not stain the liver tumors, the tumor RNases must have been derived from cells of other than parenchymal origin; this indirectly supports the results of the histochemical studies referred to above. Because of the critical role of the nucleic acids in transmission of genetic information and in protein (and enzyme) synthesis, and the

LOCALIZATION OF NUCLEASES

413

value of the enzymatic approach in studies on the biochemistry of cancer ( 2 4 1 ) , it is t o be expected that further investigations on nucleolytic enzymes in carcinogenesis will be actively pursued. Undoubtedly there is an important requirement for more standardized quantitative methods. But i t may be profitable to place increased emphasis on the specificities of the enzymes involved. It would be of interest to know, e.g., whether leukemic cell phosphodiesterase (Section 11) is a specific tumor enzyme. The problems involved in such studies have been clearly set forth by Bergel (233) and Potter (241),among others. Various attempts have been made to control tumor growth by direct hydrolysis of nucleic acids, e.g., hy daily injections of high doses of RNase, apparently causing a transient regression of solid tumors in mice (24.2).Similar transient regressions provoked by RNase and DNase have been reviewed by Bergel (233), but there is no real evidence to support the claims that hydrolysis of tumor nucleic acids is really achieved. Both RNase and DNase have been shown t o be capable of penetrating membranes of mammalian cells (243, 244), with widely varying effects. I n any event, the foregoing has in part stimulated studies on intracellular levels of nucleolytic activity accompanying tumor regression induced by various agents, and led to the discovery of elevated RNase activity associated in some cases with regression of tumor growth. For example, regression of lymphosarcoma under corticosteroid treatment is accompaiiicd by a twofold increase in cellular acid RNase activity; this increased activity was shown to arise from tumor cells and not from the macrophages associated with them (245). Treatment of lymphosarcoma with 9-a-fluoroprednisolone also leads t o increases in acid RNase activity, but this occurs prior to tumor regression, with no modification of the intracellular distribution of the enzyme (246). It should be recalled, in this connection, that fluoroprednisolone similarly affects the nuclease activities of normal lymphoid tissues (see above). I n tumor strains resistant to steroid treatment but sensitive to 5-fluorouracil, regression caused by the latter is not accompanied by an increase in RNase activity. Mashburn and Wriston (247) have similarly reported on changes in RNase activity preceding regression of lymphosarcoma in strains sensitive to a-asparaginase. Treatment with N,N'-diethylene-N'-phencthylphosphoramide of Walker 256 rat tumors reduces the rate of tumor growth, and this is accompanied by a reduction in intracellular RNA and an increase in alkaline RNase (248). A variety of cytostatic agents that limit the proliferation of ascites tumor cells provoke a correlated increase in free and latent alkaline RNase activity (249). These observations, together with additional findings (226), are interpreted as indicating that some agents, provoking regression, inactivate

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DAVID SHUGAR AND HALINA SIERAKOWSKA

tissue inhibitors, leading to increased nucleasc activity. I n the case of acid RNases this may be the result of rupture of lysosomal membranes, the liberated acid RNase then participating in autolytic processes. It nonetheless remains to be established whether nuclease activation provokes regression, or whether it is one of the side effects accompanying such regression.

IX. Possible Functions of Nucleolytic Enzymes It is generally agrecd that, in mammalian organisms, nucleases fulfill a variety of functions. Some of them are known to be involved in the digestive processes, e.g., the intracellular digestion of food in the alimentary tract by nucleases of the pancreatic juice and saliva, which is reasonably well understood. The localization of RNases, DNases and phosphodiesterases in the serous cells of digestive glands (114, 122, 184, 191, 192) is in accord with this. Intracellular digestive functions are carried out by acid nucleases localized, along with other acid hydrolases, in the lysosomes ( 1 4 4 ) . Under normal conditions, the lysosomal enzymes hydrolyze intravessicularly materials rejected by, or foreign to, the cells. It is believed that thc compartmentalization of these enzymes by the lysosomal membrane serves to regulate their activity, thus rendering them innocuous toward cellular nucleic acids. I n agreement with this is the absence of any known natural specific inhibitors of acid RNases and the lack of activity toward these enzymes of alkaline RNase inhibitors. The structure-linked latency of acid nucleases is modified as a result of cellular injury or tissue regression. Rupture of lysosomes is accompanied by the appearance of elevated levels of acid nucleases in the supernatant. These nucleases are generally presumed to be irivolvcd in autolytic processes. The occurrence of RNase (250-253) and DNase (251-253) activities on the surface of mammalian skin in somewhat baffling from the point of view of possible function. Studies on interaction with viral RNA have also shown that HeLa (254) and normal mammalian (255) cells exhibit surface-associated RNase activity, the significance of which remains to be clarified (see below). It is also generally accepted that nucleases participate in some manner in the intracellular metabolism of genetically and informationally active nucleic acids. This subject has received a good deal of attention in recent years, particularly with regard to enzymes active against RNA. Studies in this field have been confined mainly to bacteria; but, from the general similarities in basic metabolic pathways, and the results of some

LOCALIZATION OF NUCLEASES

415

studies on higher organisms, it may be inferred that the results obtained with microorganisms are also applicable to higher organisms. A tentative hypothesis for the intracellular role of ribonucleases was advanced by Elson (256, 257) following the discovery of latent ribosomal nuclease activity in E . coli. The enzyme initially released 2’:3’phosphate-terminated oligonucleotides and then hydrolyzed the cyclic phosphate rings with a preference toward cytidine and adenosine linkages ( 2 5 8 ) . Elson postulated that the latent ribosomal enzyme, which constitutes the total RNase activity of the cells, might be involved in removal of mRNA from the ribosomal surfaces. This hypothesis received a major set-back following the discovery by Neu and Heppel (105, 259, 260) that latent ribosomal RNase could be released from E . coli cells without destruction of their viability or the integrity of their ribosomal population. Formation of E . coli spheroplasts Icd to the re!ease of a large fraction of the RNase activity, with the simultaneous disappearance of a corresponding amount of latent RNase (with nearly identical properties) from the ribosomes. It was further shown that free E . coli RNase (or pancreatic RNase) was readily adsorbed and “masked” by ribosomes, but was released on breakdown of the latter. E. coli ribosomes were able to bind a 26-fold excess of the RNase activity normally associated with them upon isolation (261). RNase activity similar to that released during formation of spheroplasts was recently found in the debris of E . coli ( 2 6 2 ) . This has raised serious doubts as to whether LL1atent’’RNase is localized on the ribosomes, rather than in the vicinity of the cell wall along with other degradative enzymes, being adsorbed and masked by the ribosomes during disruption of the cell structure. I n conjunction with supernatant inhibitors, this would account for the stability of polysomal structures in the intact cell (263, 264). Reservations have also been expressed as to whether the RNase of E . coli is responsible for inactivation or degradation of mRNA. Hydrolysis of the latter by ribosome-associated enzymes leads to formation of nucleoside 5’-mono- and -pyrophosphates; no fragments with terminal 3’-phosphate groups are found among the products (107, 265). Formation of 5’phosphates is caused by a K+-activated phosphodiesterase (265, 266) ; 5’-pyrophosphate products, and the activating effect of phosphate ions, are indicative of the action of polynucleotide phosphorylase (265, 267, 268). Only when the Mg” conrtntration is lowered, leading to ribosome dissociation, do nucleosi(1e 3’-phosphatcs appear, because of activated RNasc (269). Furthermore, E . coli spheroplasts, devoid of RNase, have bccn shown readily to degrade rapidly labeled RNA ( 2 7 0 ) . Escherichia coli RNase-

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deficient mutants, with less than 1% of the RNase activity of the wild type, produce rapidly labeled RNA with a half-life identical with that of tlic wild type. Extracts from these cells support mRNA- or poly Ustimulated in vitro protein synthesis as effectively as the RNase-positive strain ( 2 6 1 ) . E . coli mutants deficient in RNase I and in both RNasc I and polynucleotide phosphorylase were shown to inactivate mRNA of induced P-galactosidase a t a rate equal to that for the wild strain (270a). Degradation of mRNA in RNase-deficient mutants of E . coli yields nucleoside 5’-mono- and -pyrophosphates ( 2 7 1 ) .Alcaligenes faecalis with no detectable RNase activity degrades both mRNA and rRNA via polynucleotide phosphorylase and K-activated phosphodiestcrasc (272).Singer and Tolbert (273) found no change in the level of 5‘-phosphate-forming, K+-activated RNase in strains dcficient in latent RNase ; in particular, they established that the enzyme, partially bound to ribosomes, is specific for polyribonucleotides in the random coil configuration. It consequently appears that the in vivo inactivation and degradation of mRNA is not due to formation of products with terminal 3‘-phosphatcs by RNase action, but is regulated in lower organisms by phosphodiesterases forming nucleoside 5’-phosphates and by polynucleotide phosphorylase, albeit the evidence regarding the role of the latter enzyme is still somewhat contradictory. These ribosome-associated enzymes degrade mRNA niorc readily than either tRNA or rRNA (107, 274). Latent RNase, on the other hand, hydrolyzes tRNA morc readily than other types of RNA ( 2 6 0 ) . Intact E . coli ribosomes are relatively resistant to endogenous latent and exogenous pancrcatic RNase a t Mg‘+ concentrations above 5 m M ; lower Mg2+ concentrations lead to their dissociation to smaller units, which are susceptible to the RNases ( 2 7 5 ) . I n intact E . coli cells subjected to phosphorus deprivation, initial degradation of rRNA to 3’-phosphate-terminated oligonucleotides is due largely to latent ribosomal RNase ( 2 7 6 ) . The probable localization of latent E . coli RNase in the region adjoining the cell membrane, together with its nonindispensibility for cell viability, imply some extracellular function. The surface localization of a number of E . coli degradative enzymes (phosphatases and nucleases) (277) is in agreement with such a concept. It is of interest in this connection that metazoan (255) and HeLa (254) cells contain a surface-associated RNase activity that appreciably decreases the infectivity, and leads to fragmentation, of infectious viral RNA. Thc existence of inicroorganims deficient in polynucleotide phosphorylwe points to the nonindispensibility of this enzyme as well. Kimhi and Littauer (278) found less than 10% of polynucleotide phos-

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phorylase firmly bound to washed ribosomes of E . coli. The specific activity of polysomes was low i n the heavy polysome region, and increased toward the smaller polysomes; such a distribution is in accord with that expected for activity bound to mRNA. Elson’s original work stimulated a series of investigations on latent ribosomal RNase in mammalian cells (46, 1/31),particularly with respect to the stability of mRNA (279, 280). Calf pancreas ribosomes were found to contain considerable RNase that differed from that of liver ribosomes in that it was precipitated by serum antibodies to crystalline pancreatic RNase. The ribosome-bound RNase was shown to exchange readily with added labeled paiicreatic RNase (281). Recently Utsonoiniya and Roth (166) found the alkaline RNase of rat liver polysomal preparations to be associated with subunits or degradation products of the ribosomes; intact ribosomes or polysomes were devoid of latent RNase and exhibited no masking of small quantities of exogenous pancreatic RNase or supernatant RNase, which associated with the smaller subunits. I n the presence of spermine or Mg”, these subunits re-form larger units, with concomitant masking of their RNase (9). On the other hand, treatment of guinea pig pancreatic ribosomes with spermine provoked a release of RNase (282). Of particular interest was the discovery of 5’-phosphate-forming endoribonuclease and phosphodiesterase I in the membranes of the endoplasmic reticulum of rat liver cells (46) and Ehrlicli tumor cells (44). Since protein synthesis was found to be more active in ribosomes attached to membranes (683),the inference was drawn that these enzymes may play some significant role in the protein synthesis machinery ( 4 6 ) . However, the possible role, if any, of polynucleotide phosphorylase in the degradation of mRNA of higher organisms is not clear. Hymer and Kuff (284) found the initial breakdown of high-molecular-weight rapidly labeled RNA of plasma cell tumor nuclei to 4-6 S components to be mediated by an enzyme resembling the supernatant ribonuclease. Harris (285) observed that the rapidly labeled RNA of HeLa cell nuclei is degraded to acid-soluble fragments by polynucleotide phosphorylase. The degradation of rRNA in rat liver microsomes has been claimed to be due to EDTA-stimulated RNase and a phosphodiesterase (286). The potential role of RNase in the regulation of mRNA-dependent protein synthesis has comparatively recently been advanced as a conceivable interpretation for the finding that maize endosperm, homozygous for the opaque-2-gene, exhibits a multifold increase in RNase activity paralleling a reduction in zein synthesis and modification of the amino acid composition of the endosperm (286a, 286b). The increase in RNase activity is apparently not due to removal of an inhibitor, and it was

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proposed that the opaque-2 locus may be a rcgulator of RNase synthesis (286~). Insofar as the DNases are conccrncd, there is now reasonable evidence pointing to some regulatory role for this class of enzymes in DNA synthesis. An increase in thc priming effectiveness of DNA by endonucleases releasing 3’-hydroxyl-terminal products has been observed with DNase I (287) and E. coli endonuclease I (288). While as a rule, DNA priming activity is reduced by endonucleases that produce 3‘phosphate-terminal products (289), at least one instance is known where the simultaneous presence of a specific DNA 3‘-phosphatase exonuclease not only removes this inhibition but results in additional enhancement of priming activity (288). These findings are further substantiated by the more recent observations of Buttin and Kornberg (290), who showed that E . coli endonuclease may play a key role in priming in vivo DNA synthesis under specific conditions. Attempts have been made to correlate activation of DNA polymerase activity by DNase I with high activity of the latter in young animals engaged in active DNA synthesis (291), but these findings are untenable in the light of further evidence (251, 291). Nucleases in Cellular Repair Mechanisms. Various microorganisms and mammalian cells are known to undergo recovery in thc dark following ultraviolet and ionizing radiation damage (292, 293). This process is, in the case of UV, accompanied by the release of small oligonucleotides containing pyrimidine dimers. The removal of these damaged fragments, which are a block to DNA synthesis and cell division, must be by means of some endonuclease(s) and is followed by repair (or “patching”) of the damaged (or “excised”) region by cellular polymerases. The significance of these “excision” endonucleases is further enhanced by the fact that other types of lesions in DNA may also be excised by them [see, e.g., reference ( 2 9 4 ) ] .I n a t least one instance, a crude cellular extract, from Micrococcus lysodeilcticus, was shown to act as an endonuclease relatively specific for DNA containing radiationinduced lesions (295, 296). It is, of course, conceivable that some nucleases involved in the excision of radiation-induced or other lesions may be known enzymes, for example, a DNase active preferentially against denatured DNA, which would be capable of excising the lesions because of the accompanying loss of secondary structure in the damaged region. But there now remains little doubt as to the existence of specific excision endonucleases, which form an integral part of some cellular machinery whose function it is to reverse or repair damage to essential genetic elements (293). An additional example of this class of enzymes is a nuclease in Bacillus subtilis that inserts breaks in DNA treated with the mono-

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functional alkylating agent, methyl methanesulfonate (MMS) . The enzyme is quite inactive against DNA extracted from cells that have recovered from MMS damage. Since the recovery process is accompanied by thymidine incorporation, it probably proceeds via excision and repair in a manner similar to, but not necessarily identical with, that for UV repair ( 2 9 7 ) . In a t least one instance, the excision process has been demonstrated in witro, the substrate being UV-irradiated DNA (i.e., DNA containing pyrimidine dimers) and the source of enzyme an extract from MicroCOCCUS lysodeikticus (298). It is to be anticipated that the purification of some such enzyme and a delineation of its properties will enhance our understanding of the mechanisms of mutagenesis and the capacity of the living cell for dealing with them. ADDENDUM The a-naphthyl derivatives of pyrimidine nucleotides have been applied to the cytochemical localization of alkaline RNase and PDase I in the central nervous system of the rat ( N I ) . l The localization patterns for both enzymes were similar, with most intense activity in the endothelium of blood vessels, the leptomeninges and membrana limitans superficialis of the brain, and the ependymal cells lining the surface of the ventricules, and the canalis centralis of the spinal cord. No activity was observed in the neuronal and glial cells. Initial trials in our laboratories have led to the preparation of the a-naphthyl esters of two purine nucleotides, 3’-IMP and 3’-AMP. The former was obtained by treatment of 2 ,5 -di-O-tetrahydropyranylinosine with a-naphthylphosphate in the presence of DCC; the latter by treatment of NG-acetyl-2 ,5 -di-O-tetrahydropyranyladenosine(obtained by enzymatic dephosphorylation of NG-acetyl-2 ,5 -di-O-tetrahydropyranyladenosine 3’-phosphate) with a-naphthylphosphate in the presence of DCC. Both of them were found to be rapidly hydrolyzed by RNase T,, while preliminary trials demonstrated their relative resistance to RNase T, and, of course, to pancreatic RNase. The stability of both substrates was sufficiently satisfactory for use in both section cytochemistry and colorimetric assays of the appropriate enzymes. Preparative methods for these substrates are therefore being elaborated. These findings considerably extend the range of application of the azo-dye coupling techniques to localization of nucleolytic enzymes. [The reader is referred to the Note Added in Proof, p. 427.1 ACKNOWLEDGMENTS

Ke are indebted to the following for making available photographs and manuscripts prior to publication: Drs. R. Daoust, G . delamirande, B. Ehinger, U. Z. ‘See reference list on p. 429.

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Littaucr, J. S. Roth, T. R. Shanta., M. Laskowski, Sr., D. Elson, A. Michalowski, and J. Jasinska; and to the Wellcome Trust and the World Health Organization for support.

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NOTEADDEDIN PROOF Several new and rather interesting nucleolytic enzymes have been described since the completion of this review. An enzyme that specifically hydrolyzes phosphodiester linkages in RNA and synthetic polynucleotides containing a 2’-O-methyl in the ribose moiety has been isolated from Amcystis nidulans ( N 2 ) . The enzyme, referred to by the authors as 2’-O-methyl RNase, gives rise to products with 5’-phosphate terminal groups. Its specificity is testified to by the fact that it does not hydrolyze DNA and is not appreciably active against normal RNA or bis-p-nitrophenyl phosphate or p-nitrophenyl 5’-TMP. It is fairly heat stable and optimally active a t about pH 7.5. No such activity has been reported in higher organisms, but the remarkable specificity of this enzyme, and its potential importance in sequence studies, will undoubtedly stimulate further studies on its distribution and properties. Ehrlich ascites tumor cell nuclei have been found to contain an exoribonuclease active a t pH 7.69.2, with a preference for single-stranded, random coil RNA which is hydrolyzed to 5’-mononucleotides ( N S ) . The enzyme is inactive against DNA and p-nitrophenyl 5’-TMP. A similar enzyme is localized in rat liver cell nuclei. An acid DNase has been isolated from malignant tumors of C3H mice and partially purified ( N 4 ) .It is an endonuclease with a preference for double-stranded DNA, which is hydrolyzed to products with 3’phosphate terminal groups. It differs from splenic DNase I1 in its preference for a d(ApT) linkage.

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Further progress has been reported in the purification of DNase I inhibitor from rat serum ( N 5 ) . The inhibitor is protein-like in nature and acts by stoichiometric binding to the enzyme. It is inactive against DNase I1 and DNA polymerase. It is also of interest that rat bone marrow, which contains mainly mid RNase and relatively little of the alkaline enzyme, has been shown to be devoid of alkaline RNase inhibitor ( N 6 ) . Additional useful reviews have appeared on DNases ( N 7 ) , on ribosomal enzymes, including RNases ( N 8 ) , and on the properties and assay of tissue nucleases ( N 9 ) . The latter adduces further data on the specificity of acid RNase, which is postulated as being specific for the internucleotide linkage between a pyrimidine nucleoside 3’-phosphate and the adjacent nucleoside. Previously reported hydrolysis of other bonds is ascribed to contamination with alkaline RNase and/or spleen PDase.2 The author also proposes a survey procedure for nuclease assays in homogenates, essentially as follows: One half the homogenate is treated with acid to inactivate RNase inhibitors, 5’-RNase and PDases I and 11, and then assayed for acid and alkaline 3’-phosphate forming RNases; the other half is used to assay 5’-RNase against poly-A as substrate, and PDases I and I1 against p-nitrophenyl 5’-TMP and 3 TMP, respectively. It should be noted that the use of the p-nitrophenyl nucleotide esters for assay of PDases is to he strongly recommended, in place of the still widely employed bis-p-nitrophenylphosphate, for reasons other than merely greater specificity and higher rates of hydrolysis. The latter compound has been found to be an active inhibitor of carboxylesterases ( N 1 0 ) . It turns out that the inhibitory effect is due to phosphorylation of the carboxylesterase molecule with concomitant release of two moles of p-nitrophenol. Consequently, release of p-nitrophenol on incubation of the substrate with tissue homogenates may, in some instances, be due in whole or in part to the presence of carboxylesterase rather than PDases, particularly since the inhibition effect is rapid a t p H 8. An extremely informative and succinct review ( N 1 1 ) discusses the selective release of E. coli degradative enzymes by successive EDTA and “shock” treatment, and summarizes the available evidence for the presumed surface localization of these enzymes, i.e., in a region between the cell wall and the cytoplasmic membrane. However, the same article contains an addendum which points to the weakness of some of the arguments and emphasizes the need for additional evidence. As regards the still controversial problem of RNases in tumor cells, it has been reported that Ehrlich ascites tumor cells contain acid PDases = phoephodiesterases.

429

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RNase but exhibit no alkaline activity, either in the presence or absence of p-chloromercuribenzoate ( N l d ) . Alkaline RNase was found only in cell suspensions contaminated with blood, and it therefore appears that it is derived from the host tissues, via the blood, in the later stages of tumor growth. Further progress has been achieved in isolation of the repair enzyme found in the wild type strain of Micrococcus lysodeikticus (see p. 419). Column chromatography led to the clear separation of an enzymatic coniponent with endonucleolytic activity against UV-irradiated D N A and with no activity against normal DNA (N1.9). The enzyme exhibited no requiremcnt for Mg2+. Somewhat surprisingly, an eytract from a mutant strain with similar UV sensitivity was only slightly active against DNA. Numerous attempts have been made to determine whether the enzymatic genetic repair mechanisms t h a t exist in microorganisms are also to be found in mammalian cclls. No such direct evidence has yet been forthcoming. But Rauth ( N l 4 ) has noted some striking similarities between dark reactivation of UV damage in mouse L cells and that in hacteria. It is to be expected that this subject will continue to elicit considerable interest, the more so in t h a t it had been previously shown, as might be expected, t h a t thymine dimers are induced by UV irradiation in the chromosomal D N A of Chinese hamster cells ( N 1 4 ) .

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