Multiple RNA polymerase species from rat liver tissue: Possible existence of a cytoplasmic enzyme

ARCHIVES

OF

BIOCHEMISTRY

Multiple

RNA

AND

Polymerase Existence

K. H. SEIFART, Institut

fiir

161, 619-632 (1972)

BIOPHYSICS

Species

from

of a Cytoplasmic B. J. BENECKE

Chemie, 355 MarburglLahn,

Physiologische

Received

Rat Liver

January

Tissue:

Possible

Enzyme AND

P. P. JUHASZ

Lahnberge,

21, 1972; accepted

Bundesrepublik

Deutschland

May 8, 1972

DNA-dependent RNA polymerase species have been extracted and purified from rat liver tissue. Apart from the established nuclear forms of the enzyme A (or I) and B (or II), an RNA polymerase activity has been purified from the cytosol of the liver cell which has been designated tentatively as C. The observed activity requires the nucleoside triphosphates ATP, GTP, CTP, and UTP, and a DNA template which cannot be substituted by synthetic ribopolymers or RNA. The incorporation of UTP catalyzed by enzyme C is resistent to rifampicin but can be suppressed by actinomytin D, and taken together with the high molecular weight of the enzyme, it is very unlikely that it belongs to a polynucleotide phosphorylase or terminal nucleotidyl transferase activity, elongating preexisting RNA chains. It can be shown that enzyme C does not emanate from a mitochondrial source and is different from both nuclear enzymes A and B by a number of suitable criteria. Phosphocellulose chromatography differentiates enzymes A and C, and chromatography on DEAE-cellulose and hydroxylapatite columns show B and C to be clearly different. In addition, functional tests show that the enzyme found in the cytoplasm is not identical to either of the nuclear enzymes A and B. Titration curves with the specific inhibitor or-amanitin show that the enzyme obtained from the cytosol is basically sensitive to amanitin, although the required concentrations of inhibitor exceed those for the B enzyme by two to three orders of magnitude. This amanitin sensitivity of enzyme C, which is intermediate between that observed for enzymes A and B, seems to clearly differentiate these three components. This observation is substantiated by other functional tests like ionic and template dependence, the capacity of catalyze pyrophosphorolysis, and t,he ability of a protein factor (23) to stimulate transcription by enzymes A, B, and C.

DNA-dependent RNA polymerase (EC 2.7.7.6.) has in recent years been isolated from a number of mammalian and other eucaryotic tissues (for a good summary see Ref. (1). A general phenomenon unifying most of these rather diversified tissues is the fact that in almost all cases this enzyme has been found in multiple forms. These forms have initially been designated by Roeder and Rutter (2) as I, II, and III for the RNA polymerases from rat liver and sea urchin according to their elution order from DEAESephadex, and as A and B by Chambon and his collaborators for the calf thymus enzymes according to their sensitivity to Qamanitin. This toxin from the toadstool amanita phalloides inhibits nuclear RNA

synthesis as was initially shown by Stirpe and associates (3). This inhibition is a specific phenomenon involving polymerase B (or II) as was initially shown for rat liver by Seifart and Sekeris (4) and calf thymus by Kedinger et al. (5). These studies have since been confirmed and extended to a large number of other cases (6-9). Recently it has been shown (10-12) that the B enzymes from calf thymus and rat liver consist of two subspecies, B, and B,, , and also the A enzymes from a number of tissues seem to occur in several subclasses (13-15). It is currently accepted that enzyme A (or I) is structurally associated with the nucleolus and is chromatographically distinct. and functionally different,, with respect

519 Copyright All rights

@ 1972 by Academic Press, of reproduction in any form

Inc. reserved.

520

HEIFART,

BENECKE,

to template and ionic requirements for optimal synthesis, from enzyme B (or II) which is structurally located in the nucleoplasm. It is generally agreed upon that all t’hese enzymes described are located in the cell nucleus.

This report describes the purification of nuclear enzymes A and B from rat liver and contra&s these enzymes to an RNA polymerase activity observed in the caytjosol of this tissue, which has tentatively been called C. This enzyme has been purified to a high degree and may represent an entity distinct from the nuclear or mitochondrial RNA polymerases. MATERIALS

AND

METHODS

Materials. a-Amanitin was a generous gift by Prof. Th. Wieland (Heidelberg). Rifampicin and derivatives were kindly donated by Dr. L. Silvestri Lepetit spA (Milano). [3H]UTP (1-5 Ci/ mmole) was obtained from the Radiochemical Centre (Amersham) ; nucleosidetriphosphates, creatinphosphate, creatinphosphokinase from Boehringer (Mannheim) ; DEAE- (DE32,l meq/g) and phosphocellulose (P-11, 7 meq/g) from Whatman (London) ; hydroxylapatite from Cla.rkson Chem. Co. (Williamsport, NY); Sephadex G-75 from Pharmacia (Uppsala); synthetic ribopolymers, actinomycin D, polyethyleneglycol and materials for polyacrylamide gel electrophoresis from Serva (Heidelberg), and salmon sperm DNA (highly polymerized, A grade) from Calbiothem (Los Angeles). All other chemicals were reagent grade from Merck (Darmstadt). Buffers employed were: TSS-buffer (Tris-Cl, 0.05 M, pH 7.5, 0.25 M sucrose, 0.010 M MgClz, 0.025 M KCl). Buffer I: (TSS, but containing 2.2 M sucrose). Buffer II: (Tris-Cl, 0.05 M, pH 7.9, 0.25 mM EDTA, 5 mM fl-mercaptoethanol, 20% glycerol). Buffer III: (Buffer II + 1.0 M (NHJ2SO,). Buffer IV: (0.02 M phosphate, pH 6.8, 0.25 mM EDTA, 5 mM p-mercaptoethanol, 20% glycerol). Mehods. RNA polymerase activity was measured in an in vitro system, incubated for 40 min at 37”C, consisting of 0.2 pmoles each of ATP, GTP, CTP; 0.002 rmoles UTP, 1.0 FCi [3H]UTP, 1.5 rmoles creatinphosphate, 5 pg creatinphosphokinase, 1.5 rmoles p-mercaptoethanol, 10 pmoles Tris-Cl, pH 7.9, 20 pg native DNA, 0.5 pmoles MnSOl , 10 pmoles (NH,)xSOa (resulting ionic strength = 0.2), and variable amounts of RNA polymerase in a final volume of 150 ~1. At the end of the incubation, an aliquot was pipetted onto filter paper discs (16 cm2, Schleicher and

AND

JUHASZ

Schuell 2043 b), precipitated, and washed five times in 1 liter each of ice cold 5% perchloric acid, washed with ethanol and diethyl ether, and counted by liquid scintillation in toluene (5 g PPO, 200 mg POPOP/liter). The extraction and purification procedure employed to obtain the RNA polymerase species has been partly modified from previously pub-

lished methods (2, 5, 16) and is summarized in Fig. 1. All operations were carried out at 2-4°C unless otherwise stated. Liver nuclei were pre-

pared from 120. to 150-g male Wistar BIZ. II rats by a modified procedure of Chauveau et al. (17). Livers were flushed in situ with 10 ml ice cold TSS-buffer, minced with scissors followed by glass-Teflon homogenization. Typically 5 g liver were homogenized in about 30 ml TSS. After centrifugation for 7 min at 1OoOg the supernatant’

fluid was retained and the pellet containing nuclei

was suspended

in approximately

buffer I. The cytoplasmic

supernatant

the

10 vol

fluid was

centrifuged for 30 min at 48,000g (max) and the postmitochondrial supernatant fluid was ultracentrifuged at 180,OOOg (av) for 1 hr. After aspiration of flotated lipids, this cytoplasmic supernatant fluid was used as source of one of the polymerase species. The nuclear suspension in buffer I was layered over a lo-ml cushion of the same buffer in each of 24 tubes and centrifuged for 2 hr at 95,OOOg (max) in the R 30 rotor of the Beckman centrifuge. These purified nuclei were employed to isolate nucleoli by the method of Busch and co-workers (18) or were used directly for the extraction of RNA polymerase. In a typical experiment nuclei from 500 g liver CRUX

! PHOSPbO-CELLULOSE

DENSITY

I GRClD,EbTS

FIG. 1. Schematic dure RNA

RAT il”ER

‘IOMOGENAIE

I PC 1 HVOROXYLAPATITE

0

! GRADIENTS

I PC HAP I 0 GRADIENTS

representation of the proceemployed to extract and purify multiple polymerase species from rat liver tissue.

RNA

POLYMERASE

SPECIES

tissue were suspended in 700 ml extraction-buffer III and stirred for 1.5 hr at 2°C. Proteins were subsequently precipitated from the medium by the addition of crystalline (NH,)zSO, (33 g/100 ml), a procedure which maintains the bulk of the DNA in solution. Sedimented proteins were suspended in approximately 150 ml buffer II and dialyzed against the same buffer overnight. After discarding resulting precipitates by centrifugation at 12,000g (max) for 15 min the supernatant fluid was applied to a DEAE-cellulose column which was then extensively washed and subsequently developed according to the details described in Fig. 2. This procedure separates the total nuclear RNA polymerase activity into two main components A and B by the nomenclature of Chambon and co-workers. Simultaneously, approximately 500 ml of the cytoplasmic supernatant fluid was applied to a separate DEAE-cellulose column after the material had been adjusted to pH 7.9, 200% glycerol and a conductivity of max 1.5 mS by the addition of buffer II. Details of the chromatography are as described in Fig. 2. The enzymatic activity recovered under each of the DEAE-cellulose peaks A, B, and C was pooled, dialyzed for 4 hr against 10 vol of buffer B, and rechromatographed separately on three individual phosphocellulose columns according to the details described in Fig. 4. The enzymatic activity recovered under peaks B and C on phosphocellulose was pooled separately, dialyzed against 10 vol of phosphate buffer IV, and applied to two identical hydroxylapatite columns. The enzyme activities B and C, eluted from these columns as described in Fig. 5, as well as the A-activity eluted from the phosphocellulose column, was concentrated by dialysis against 30% polyethylene glycol in buffer IT and centrifuged through density gradients as described in Fig. 6. General methods. Electrophoresis of the peak fractions was conducted through 4% polyacrylamide gels containing SDS according to the method of Weber and Osborn (19). Binding of radioactive [‘*Cl-methyl-r-amanitin to enzyme fractions A, B, and C obtained from either DEAEor phosphocellulose was studied by incubation of these fractions with an excess of the labeled inhibitor for 20 min at 37°C. Controls were conducted by incubating an identical RNA polymerase fraction with buffer in the same manner. Separation of free and protein-bound amanitin was achieved by gel-filtrating 1.0 ml of the incubate on a 1 X 30-cm Sephadex G 75 column equilibrated in buffer II, and counting a 250-J aliquot of each fraction in 10.0 ml Bray’s scintillation fluid (20). Concurrently RNA polymerase activity was assessed in the fractions of all three (A, B, and C) amanitin vs control sets. Protein

FROM

RAT

LIVER

521

was measured by precipitation in 1.1. M TCA and subsequent recording at 400 nm by the method of Luck et al. (21). RESULTS

RNA polymerase activity, obtained by extraction of purified nuclei in the fashion described, can be separated into two components A and B on DEAE-cellulose as has also been described by several other workers (1). As is shown in Fig. 2 (Part II), these components elute at ionic strengths of 0.2 (A) and 0.3 (B), respectively, with considerable purification. Enzymatically inactive protein which is not bound to DEAE-cellulose, and is consequently excluded from the column, is not shown in Fig. 2. If RNA polymerase is ext’racted from prepurified nucleoli by the same technique described above for the nuclear enzymes, DEAE-cellulose chromatography (Fig. 3) reveals a single peak of activity, eluting at the same ionic strength found for nuclear enzyme A (0.20-0.21), indicating t)hat only this enzyme is associated with the nucleolus, although it is not clear whether all of the nuclear A enzyme is in fact associated with this organelle. Chromatography on DEAE-cellulose of the ultracentrifuged cytosol (Fig. 2, Part I) obtained by the method described, reveals a single peak of activity (designated as C) eluting at an ionic strength of about 0.22 which, for all practical purposes, is indiscernible from that observed for nuclear polymerase A on this exchanger. No activity is found in the chromatographic region of nuclear polymerasc B. Chromatography of activities A, B, and C on phosphocellulose (Fig. 4) reveals an elution order of C:B:A. The relationship of B to A on this exchanger suggests a more acidic nature of B already found on DEAEcellulose. This classification is not as clear for enzyme C which elutes before enzyme A on phosphocelullose but approximately equiionic from DEAE-cellulose. The rather slight difference in elution point’s between B and C on phosphocellulose (Fig. 4) can be substantiated on hydroxylapatite by a definitely different elution patt,crn (Fig. 5). The latter exchanger,

522

---. ~__I r SEIFART,

7300 I 2250 E '200, ii F I50

BENECKE,

AND

JUHASZ

!I

45

30

15

FRACTION

NR.

60

75

FIG. 2. 1)EA.Scellulose chromatography of the cytosol on a 4 X l&cm column (part I) and the nuclear extract on a 2.5 X 13-cm column (part II). Elution of approximately 8.0.ml fractions was achieved with a linear gradient (500 + 500 ml; 0.0-0.3 M NH&l in buffer II) pumped from an ultrograd mixer at a flow-rate of about 60 ml/hr. I -.

-1

I i 9

1

I I 0\

FIG. 3. DEAE-cellulose chromatography on a 1 X g-cm column of RNA polymerase activity extracted from purified nucleoli; a8 described in Methods. Elution of 2.5-ml fractions was achieved with a linear gradient (40 + 40 ml; 0.0-0.3 M NH&l in buffer II), at a flowrate of 10 ml/hr.

which permits significant purification of enzymes B and C unfortunately cannot be employed for enzyme A since it has not been possible thus far to recover the activity from

hydroxylapatite in a sharp peak. This matrix either inactivates the enzyme or does not allow definitive desorption of enzymatically active protein, resulting in very broad and ill-defined peaks of enzyme activity. Density gradients (Fig. 6) do not resolve enzymes B and C and show both compounds with a molecular weight of approximately 500,000 daltons, while a statistical calculation of several experiments apparently reveals a somewhat lower sedimentation coefficient for A. However, this difference has not been observed in all cases and our data do not justify strong emphasis of this point. The extraction and purification procedure for enzymes A, B, and C which is summarized in Fig. 1 yields enzyme C with a high degree of purity and enzyme B which is probably homogeneous. Enzyme A is still associated with a considerable amount of contaminating protein and additional purification steps are being probed to solve this problem. Electrophoresis of enzyme B through SDS polyacrylamide gels (Fig. 7) shows a subunit pattern with molecular weight,s of 200,

RNA

POLYMERASE

SPECIES

FRACTION

FROM

RAT

LIVER

NR.

FIG. 4. Simultaneous chromat,ography of enzyme activities A, B, and C (as obtained from DEAE-cellulose) on identical 2.5 X IO-cm phosphqcellulose columns. Elution of approximately 7.0.ml fractions was achieved with a linear gradient (350 + 350 ml from 0.0-0.6 M NH&l in buffer II) pumped from an ultrograd mixer at a flow rate of about 30 ml/hr. In order to achieve identical conditions, identical columns were packed from the same batch of exchanger and were developed at identical elution rates from one pump (with multiple outlets) from t,he same gradient mixer.

FRACTION NR

FIG. 5. Simultaneous chromatography of enzyme activities B and C on identical 2.0 X 5.5~cm hydroxylapatite columns. Elution of approximately 3.0.ml fractions was achieved with a linear gradient (50 f 50 ml from 0.0-0.4 M with respect to sodium phosphate in buffer IV) at a flowrate of about 15 ml/hr. The same precautions as described in Fig. 4 were taken to ensure maximum equality of conditions.

175, 140, 38, %2,16 X lo3 daltons judged by the subunit pattern of E. coli RNA polymerase and myosin as markers. It is almost certain that the enzyme in question represcnts a mixture of B, and B,, (10-12) which have not been separat,ed by this procedure. By the same criteria enzyme C shows t\vo high molecular-weight components which are similar to those of the B enzyme. A strict stoichiometry between high- and low-molecular weight components does not appear justified for enzyme C at present, since it is questionable whether it is completely clean and which of the low molecular-weight components actually belong to the enzyme. The important point at the moment seems to be, however, that the high molecular-weight subunit.s of B and C show some similarity although they are not identical. On the basis of these data and the chromatographic findings reported, one can conclude that the cytoplasmic enzyme is not identical to either of the nuclear enzymes A or B in all respects. However, these enzymes

524

SEIFAILT,

I

CATALASE i

u--_-

10

BENECKE,

MARKER o-o

FJ

+----+

A

20 FRACTION NR

30

I

gradient centrifugation of FIG. 6. Density enzyme activities A (after phosphocellulose, Fig. 4), B, and C (after hydroxylapatite, Fig. 5). Enzyme solutions (1.1 ml) were layered over 11.0 ml linear 5-20y0 sucrose gradients in buffer II (containing 10 instead of 20% glycerol) and cent,rifuged for 15 hr at 210,000g (max) in the SW 41 rotor of the Beckman centrifuge. Fractions (0.4 ml) were collected through a needle from the bottom of the tube.

are prepared by different procedures (Fig. 1) and are consequently associated with differing proteins which could conceivably alter chromatographic behavior. Before separation on DEAE-cellulose, nuclear enzymes A and B are extracted by a high-salt procedure, whereas this step is for obvious reasons unnecessary to obtain the act)ivity from the cytosol. In comparative studies, the cytosol was, therefore, precipitated with (NH&SO4 to mimic the high-salt step. This procedure did not, however, alter subsequent chromatographic behavior of enzyme C on DEAE-cellulose. In addition, enzymes A, B, and C were preisolated on DEAE-cellulose followed by remixing of A and C, and B and C. Subsequent rechromatography on the same exchanger shows that the elution points are faithful estimates, which are not altered by the presence of the other enzymes in the combinations tested. For this reason, the

AND JUHASZ

results have not been shown in separate figures, since they convey no additional information. Although the chromatographic differences found between enzymes A, B, and C on different sets of exchangers were indicative, functional tests were conducted with these enzymes to further elucidate this point. The catalysis of UTP incorporation into acid-precipitable material by enzyme C is completely dependent on a DNA template which cannot. be replaced by synthetic ribopolymers or RXA (Table I). Jloreover, enzymatic activity depends on the simultaneous presence of the ribonucleoside triphosphates ATP, GTP, CTP, and UTP. It could be postulated that the labeled UTP is in part dephosphorylated to UDP which could then, albeit suboptimally, be utilized by a polynucleot,ide phosphorylase type of activity. However, inclusion of UDP, 1VIg2+, and poly (U) as a primer does not stimulat’o, but in fact, totally represses synthesis (Table I). In addition, enzyme activity C is inhibitable by actinomycin D, and the chemical rifampicin derivates AF/O5 and AF/013 but is resistant to authentic rifampicin (Table II). It is not sensitive to cycloheximide as has been reported for one of the RNA polymerase species from Blastocladiella emersonii (22). Collectively, enzyme C has essentially the catalytic properties of the classical nuclear RNA polymerases A and B (Tables I and II). The action of a protein factor known to stimulate

the transcription

of enzyme

B on

native templates (23), was tested on enzymes A, B, and C. The results show, that under the conditions employed in Table II, t,his factor only stimulates RNA synthesis catalyzed by enzyme B and not that of enzymes A and C. If RNA synthesis by each of enzymes A, B, and C is measured in the presence of increasing concentrations of Lu-amanitin it bccomes clear that the sensitivity of these enzymes toward that inhibitor is completely different. Half-maximal inhibition for cnzyme B is observed at cu-amanitin concentra-

tions of approximately 0.01 pg,/ml which is in good agreement with values previously reported for this enzyme (4). In contrast, a

RNA

B

POLYMkXASE

SPECIES

E. Coli .on 4% gels

C

FROM

RAT

52.5

LIVER

E.Coli on mixed 4% gels

B and 10 %

FIG. 7. Electrophoresis through 4 y0 polyacrylamide gels containing SDS according to the method of Weber and Osborn (19). Samples, pretreated with 170 SIX3 for 2 hr at 67” and dialyzed overnight against 0.01 M phosphate buffer, pH 7.2, containing O.l’i; SI)S, were layered on 0.6 X 8.5.cm internal diameter gels. Electrophoresis was conducted toward the anode for 5 hr at 7 mA per tube, stained for 2 hr in 0.25y0 Coomasie brilliant blue, 409; methanol, 7.50/; acetic acid and destained in 20% methanol 7.5’3$ acetic acid. Gels on the right are mixed 47, and lo<;& gels (4.2 cm and 4.2 cm, respectively), to achieve an adequate demonstration of small and large subunits of enzyme B. In this case electrophoresis was conducted for 10 hr; hence the pronounced separation of the large subunits seen.

concentration of 2-3 orders of magnitude in excess of this value is required to reach the same degrecl of inhibition for enzyme C, while enzyme A is not affected at all by these concentrat’ions (Fig. 8). The presence of unspecific amanitin-binding prot’eins, shifting the titration curve and thereby the apparent K i was investigated by testing amanitin sensitivity of enzyme B in the presence or absence of enzyme C (containing such a hypothct’ically postulated compound). The results from a typical experiment show (Table III) that approximately the same degree of overall inhibition is observed for

enzyme B in either case, lending no proof t’o this hypothesis. Similar results arc conveyed by the binding studies with labeled [14C]-methyl-yamanitin (Fig. 9). These resu1t.s show quite clearly, that if enzyme C is incubat’ed with the inhibitor which is subsequent,ly removed by gel filtration, t’he enzyme activity remains clearly impaired. In addition, the bulk of t,he label appears in free form and is not protein bound. Therefore, it cannot be concluded that the lowered susceptibility of enzyme C is due merely to the presence of a strong inhibitor-binding protein since Fig. 9 shows

526

SEIFART, TABLE

BENECKE,

I

TEMPLATE AND SUBSTRATE DEPENDENCE CYTOPLASMIC RNA POLYMERASE C Conditionsa

-

-~ A. Template requirements Complete system Complete system with denatured DNA Complete system - DNA Complete system - DNA, + rRNA Complete system - DNA, +

OF

L'HIUTP incorporated into acidprecipitsble material (expressed BS % of controls)

100,O 135,o

620 290 775

~01~ A

Complete system POlY U Complete system

-

DNA, +

090

-

DNA,

+

090

-

DNA,

+

o,o

POSY G

Complete

system

POlY (=

B. Substrate requirements Complete system 100,o Complete system -GTP, -ATP 330 Complete system -CTP, -ATP 82’3 Complete system -GTP, -CTP 12,o Complete system -GTP, - ATP, 11,8 -CTP C. Test for polynucleotide phosphorylase (-DNA, +poly(U)) -Mn*, + Mg++ 2,3 (20 mM) + UDP (10 mM) conditions were as described in 5 Assay Methods. When DNA was omitted, the substituting polymer was added at the same concentration (20 pg/150 ~1. of assay medium).

the presence of huge amounts of unbound amanitin. It is interest,ing to note that enzyme B, lvhich has a very high affinity for amanitin and is completely inhibited in the presence of concentrations (50 pg/ml) used in Fig. 9, regains part of its activity if the excess of inhibitor is removed. Similar results have been reported by Chambon et al. (24) for calf thymus B using a different approach. It should be ment)ioned that in the region of enzyme A, which is at no time influenced by amanitin as shown before, a small but clear degree of radioactive binding can be registered. This can either be due to unspe-

AND

JUHASZ

cific binding of amanitin, or the inhibitor is bound to the enzyme without affecting its activity. Clarification of this interesting point must be postponed until enzyme A has been purified more extensively. By the same token it is not completely clear at present whether all the radioactivit’y recorded under the protein peaks of enzymes B and C truly reflects enzyme-bound amanitin. Since the enzymatic activity is inhibited in these cases, it is clear, however, that the radioactivity must, at least in part, reflect amanitin binding to polymerase. Functional studies concerning the ionic dependence of enzymes A, B, and C show the well-documented pattern for nuclear enzymes A and B (1) which need not be elaborated. It is interesting to note, (Fig. 10) that enzyme C, like B, optimally functions with Mn2+, but in the presence of hig2+ is depressed in its activity by added salt and in this respect resembles enzyme A. At relatively high pyrophosphate (PPi) concentrations and in t’he presence of DNA, TABLE EFFECT OF INHIBITORS PROTEIN FACTOR OF RNA POLYMERASE

Additions

Control Stimulatory Actinomycin WI a/ml) Rifampicin r&4 Rif. AF/05 a/ml ) Rif. AF/013 e/ml) Cycloheximide (400 a/ml) cu-Amanitin rg ml)

II AND

A

STIMULAT~RY

ON THE SPECIES A,

ACTIVITY

B, AND

Co

[3H]UTP incorporated into acid- precipitable material (expressed as %;i of control) Polymerase ~. ~ i\ B C

factor D

100 115 10

100 270 8

100 109 11

(167

100

100

106

(133

50

60

65

30

40

25

100

100

100

100

0

(133

(0.1

75 (see also Fig. 8)

5 Assay conditions were as described Methods. The stimulatory factor was isolated described previously (23) and was added in quantity of 30 pg protein/assay volume of 150

in as a ~1.

RNA

POLYMERASE

SPECIES

FROM

RAT

LIVER

527

RNA polymerase from E. coli is known to catalyze pyrophosphorolysis, i.e., the reverse reaction of triphosphate polymerization and elimination of PPi (25). Pyrophos-

AMANI TI N CONCENTRATION buy /ml ) FIG. 8. Effect of a-amanitin at increasing concentrations on the activity of RNA polymerase species A, B, and C. Amanitin concentrations reflect final concentrations in the assay. This result was obtained for polymerase C at all purification steps ranging from crude cytosol t)o enzyme purified over DEAE-cellulose, rechromatographed over DEAE, phosphocellulose, and hydroxylapatite.

TABLE TEST

FOR THE PRESENCE AMANITIN-BINDING

III OF UNSPECIFIC PROTEINSRNA

Conditions

Polymerase B + amanitin Polymerase C + amanitin Polymerase B + C + amanitin

polymerase activity (cpm/O.l ml) 1980 620 840 800 2740 1780

Relative amanitin inhiljition Ml) 69 5 65

o ol-Amanitin was employed at a submaximal dose of 0.03 pgsml in the assay.

FIG. 9. Binding of [‘%I-met,hyl-r-amanitin (----) to protein samples containing either RNA polymerase A, B, or C. Five microcuries amanitin was present in the incubation medium at a final concentration of 50 pg/ml. Incubation was conducted as in Methods. Separation of free and protein-bound radioactivity was achieved by gel filtration through Sephadex G-75. Concurrent,ly enzymatic activity was measured in the column fractions after filtration of control (O--O) and aminitin-treated (A--A) samples of RNA polymerase A, B, or C. Enzymes were obtained after the phosphocellulose step.

phorolysis, cat’alyzed by the three liver enzymes under investigation, was st’udied and the results are reported in lcig. 11. These data suggest that enzyme A actively catalyzes pyrophosphorolysis at I’Pi concentrations of 10 mhr. With enzymes B and C it, has been found, in contrast, that PPi concentrations of 10 rnM clearly perturb an otherwise linearly proceeding synthesis, resulting in an overall inhibition of RNA synthesis. This could be explained theoretically by a numbrr of artifactual and insignificant reasons involving factors by which the addition of pyrophosphate could unspecifically intrrfew with RXA synthesis. In the case of enzyme A, however, one observes a high dcgrcc of apparent pyrophosphorolysis of presynthesized RYA molecules (Fig. 11). Analysis of the

528

SEIFART,

Mn++

4 7

~

MO++

BENECKE,

Mg++

Mg++l

AND

JUHASZ

enzyme activity has been found in the cytosol of the liver cell which incorporates UTP into acid-pr&pitable material. Taking into account th(k terminology of Chambon and co-workers for th(> nuclear enzymes A and B from calf thymus, this RXA-polymerizing activity from thr cytosol has tentatively been calltld C. This activity has been obtained after ultraccntrifugation of wholecc41 homogenates and is not structurally

1 ,

A PP

I

.’

Pfi (l.OmM)

‘.

k. ; __~/K ‘\ , /

50

llc +

FIG. 10. Effect of Mn2+ (3.4 mM) or Mg2+ (20 mM) cations on the activity of RNA polymerase A, B, and C in the presence (67 mM) or absence of (NH4).#04. Enzymes were obtained after the phosphocellulose step. of enzyme A for possible contamination by RNAse was negative, but the possibility remains that this observation is due to nuclease attack which, theoretically, could depend (directly or indirectly) on pyrophosphate. This phenomenon is not recorded for enzymes B and C at identical pyrophosphatc concentrations, which cxteed substrate (nucleosidetriphosphatc) concentrations by at least one order of magnitude, since the K, for XTP concent,rat.ion has been calculated to bc of the order of 10-4 ar. This observation seemingly differentiates enzyme A from both B and C, and may be relat’ed to the fact’ that enzyme A synt’hesizes RNA much more readily on native DNA templates than is the case for enzymes B and C which transcribe heat-denatured DNA with differing, but clear prefpreparation

erentiality

(Ref.

23 and Table

I).

DISCUSSION

The major finding of the work reported here is that, apart from the established forms of nuclear RNA polymerases A (or I) and B (or II) (for a summary see Ref. l), an

I 5 2

c

PP. (I.0 mM) ;K

I

501 j

~~,i

:,,

?

’ Do (1OOmM)

I

251 2

FIG. 11. Pyrophosphorolysis catalyzed by enzyme fractions A, B, and C obtained after phosphocellulose chromatography. RNA synthesis in this case was conducted as in Methods with the exception that creatinephosphate and creatinephosphokinase were omitted from the incubation medium. RNA synthesis was allowed to proceed normally during the time-course indicated. At 40 min (arrow) either PPi at concentrations of 1.0 mM (O-----O) and 10.0 mM (e-.-.-O) or buffer (O---O) was added. The time course of the reaction was followed for an additional 60 min.

RNA

POLYMEKASE

SPECIES

bound to any of the established cell organelles. The synthesis of homopolyribonucleotides from ribonucleoside triphosphates has been shown to occur in a number of biological systems (26-31) and most frequently, the newly synthesized homopolyribonucleotide is added onto the end of a polynucleotide sequence required as a primer in the reaction. These reactions, which are not DNA dependent and unaffected by actinomycin D (32), can also be primed by synthetic polyribonucleotides or RNA as is the case with ATP polymerase (EC 2.7.7.19.). The incorporation of UTP into acid-precipitablc mat.erial catalyzed by enzyme C, isolated and purified as described, is completely dependent’ upon the addition of either native or denatured DNA templates and these cannot be substituted by ribohomopolymers nor the RNA species tested (Table I). The incorporating activity is as completely blocked by actinomycin D as is RNA synthesis catalyzed by any of the other nuclear RNA polymerasc types (Table II). Omission of either divalent metal cations or any one or more of the four required nucleoside triphosphates from the incubat,ion medium result,s in highly depressed incorporation values. In addition, UDP dots not support synthesis in the presence of IlIg?+ and Poly (U) as a primer (Table I). These facts, taken together with the high molecular weight of the observed enzyme activity, seem to exclude rather safely that the enzyme activity belongs to the class of a polynucleotide phosphorylase or terminal nucleotidyl t’ransferase activity, elongating preexisting ribopolymers. Initially it was suspected that the activity observed in the cytosol represents either (i) a mixture of nuclear enzymes A and B which were leached out in varying proportions during the course of homogenization, or alternatively (ii) emanates from a mitochondrial source. Mitochondrial RT\‘A polymerase (33-37) occurs in relatively rigid structural association and is not readily leached out of the organelle. This would argue against the latter possibility since the starting material, from which enzyme C is obtained, is a particlefree ultracentrifugate. Should the activity

FROM

RAT

LIVER

529

observed in the cytosol nevertheless represent mitochondrial leakage, it should be rifampicin sensitive, since RNA polymerase from rat liver mitochondria has been found t,o be inhibitable by this antibiotic (33, 34). The observed activity is, however, rifampitin resistant. In addit#ion, mitochondrial RNA polymerase from Xewospora crassa (35) and rat liver (34) has been shown to have a molecular weight of approximately 65,000 daltons, which together with the rifampicin resistance of enzyme C, renders model (ii) very unlikely. The polymerase activity is partially sensitive to the chemically modified rifampicins AI’/05 and AI?/013; this property is, however, shared by the nuclear enzymes A and B to the same extent and provides no basis for differentiation. In addition, it should be kept in mind, that a large amount of activity is found in the cytoplasm. This can hardly be explained by leakage from mitochondria which, in the rat liver, show comparatively low activities of RT\‘A polymerase (own unpublished observations). It is very difficult and in accuracy questionable to measure quantitatively RSA polymerasc activity in crude cellular homogenates due to simultaneous cont’amination by RNAse, solubility problems, inaccessibility of enzymes to templates, etc. Approximate estimates, which have nevertheless been attempted, show about, 30 % of the total activity to be located in the cytoplasm after ultracentrifugation. Similar findings although interpreted diffcrently have been reported by Chest’erton et al. (33). The former alternative of nuclear leakage initially discussed is obviously the most prevalent, one, since current preparation methods certainly damage a high proportion of nuclei and even in intact nuclei the passage of prot’eins across the nuclear membrane is a well-documented phenomenon (39). The striking observation is, however, that if the phenomenon is to be reduced to mere nuclear leakage, only one of the polymerases can be involved, since only a single peak of activity in the chromatographic region of enzyme A is recovered after DEAEcellulose chromatography of the cyt,osol (Fig. 2). By this criterion, very IMe, if any,

530

SEIFART,

BENECKE.

B activity is present in the cytosol. It should be mentioned that the crude cytoplasmic supernatant fraction before ultracentrifugation contains ample RNA polymerase activity which also bears the properties normally associated with nuclear polymerase B. This activity is, however, not DNA dependent, can be removed by ultracentrifugation, and is by all probability chromatinbound or structurally associated in some other fashion. It would be readily conceivable that polymerase A has a lesser degree of structural association and is much more prone to cytoplasmically directed diffusion, implying that the cytoplasmic polymerase would emanate from enzyme A. This would be in accord with the indiscernible chromatographic behavior of A and C on DEAE-cellulose. However, several points argue against this assumption. Simultaneous chromatography of all three enzymes on individual phosphocellulose columns under very stringent conditions seems to clearly differentiate enzyme A and C (Fig. 4). In addition, amanitin-inhibition curves (Fig. 8) show that enzyme C is, in principle, sensitive to this toxin, although very high concentrations of amanitin are required. Under identical conditions, however, enzyme A remains completely unimpaired showing that this is not a nonspecific toxic effect. Therefore, it is very unlikely that enzyme C is identical to and emanates from nuclear enzyme A. The initial classification of nuclear enzymes A and B was based on their sensitivity toward ar-amanitin (5) and on this basis one would have to classify the cytoplasmic activity as “B-like.” The preliminary subunit pattern obtained for enzymes B and C on SDS polyacrylamide gels also seems to indicate certain similarities. However, the completely different sensitivity of these two enzymes seems to justify the classification of this enzyme as C, or as a class of enzymes with intermediate sensitivity toward a-amanitin. It is interesting to note that, while nuclear enzymes B from mammalian eucaryotes are extremely sensitive to this inhibitor, comparable enzymes from the lower eucaryote yeast, for instance, are much less sensitive to a-amanitin (8).

AND JUHASZ

Titration studies of enzyme B with submaximal concentrations of amanitin in the presence of a rather crude protein fraction containing enzyme C make it very unlikely that this phenomenon can be attributed to the presence of a nonspecific amanitin-binding protein present as a contaminant. The same conclusion can be arrived at by the binding studies with labeled amanitin which clearly show that free amanitin can be removed from the protein fraction by gel filtration and that the inhibitor is by no means quantitatively bound. Moreover, this differing amanitin-sensitivity curve has been found in all purification steps ranging from the crude ultracentrifuged cytosol to the enzyme purified through hydroxylapatite. In addition, chromatographic separation of nuclear enzymes A and B show that these occur in a rough stoichiometry of 1: 1. Testing the crude nuclear extract before DEAE chromatography, or even whole nuclei, with low doses of amanitin usually reveals an inhibition estimate of approximately 50 %, indicating that nonspecific amanitin-binding proteins are apparently not present in very large quantity. Functional tests conducted with the three liver enzymes under investigation reveal certain differences, although it should be stressed that the array of meaningful functional criteria presently available is limited. Product analyses have been attempted, but have thus far not revealed any principle differences of functional significance. The divalent metal cation and ionic dependency of enzyme C has been studied as outlined and shows similarities to both A and B. Enzyme C, like B, is maximally stimulated by a Mn2+: (NH&S04 system. In the presence of RIg2+, however, added (NH&SO4 depresses synthesis; a characteristic typical for A and in contrast to B (Fig. 10). An interesting aspect is that the degree of pyrophosphorolysis catalyzed by enzyme A is much higher than that seen with B and C. It has previously been shown by work from this and other laboratories (23) that a protein factor can be isolated from rat liver cytoplasm which is similar in its action to that from calf thymus (40) and specifically stimulates the t’ranscription of enzyme B on

RNA POLYMERASE

SPECIES FROM RAT LIVER

native templates. The action of this factor has been tested on enzyme C and the results show no stimulation with enzymes A or C but markedly enhanced synthesis with B. It can, therefore, be concluded that enzyme C bears a number of functional properties usually exemplified by B, the most prevalent and conclusive one being its amanitin sensitivity, although vast differences in Ki values have been found between these two enzymes. In addition, chromatographic behavior on DEAE-cellulose and hydroxylapatite, as well as other functional tests show B and C to be nonidentical. By the same token, A and C can be differentiated both functionally and chromatographically by suitable systems. If obvious artifacts have been excluded and the existence of a cytoplasmically located RNA polymerase can be substantiated by additional work from this and other laboratories, the rather perplexing question concerning the significance of this finding and its biological relationship to the known function of the enzyme, i.e., the transcription of DNA, has to be dealt with. It is well documented that, in the case of DNA polymerase, a large portion of this enzyme is located in the cytoplasm (41-44). Attempts to differentiate the nuclear and cytoplasmic DNA polymerases have thus far not provided a conclusive answer. It has recently been reported by Weissbach et al. (45) that DNA polymerase of HeLa cells and of a normal human lung diploid line WI-38 occurs in three forms. Two of these are nuclear and different from one another; the third is cytoplasmically located and similar by a number of criteria to one of the nuclear enzymes. These findings are very similar to the results reported in this investigation, although the implications are not clear in either case. Precursor relationships or transformation phenomena between these enzymes are, of course, possible, although no, or very few data are available to solidify this concept. Chesterton and Butterworth (13) have reported multiple forms of nuclear RNA polymerase A and have discussed possible transformations. It should be mentioned, however, that with the extraction procedure employed in this investigation only one form

531

of nuclear A enzyme is obtained on DEAEcellulose which does not split into multiple components on phosphocellulose. Concepts, by which the cytosol contains informational DNA (46) which is transcribed extranuclearly, have been described. Should it be possible to solidify these hypotheses, it is conceivable that the RNA polymerase in the cytosol plays a role in this context. By current concept, the cytoplasm seems to be the site of synthesis for most if not all of the established nuclear proteins. Gallwitz and Miiller (47) initially showed this to be the case for the histones and although nuclear protein synthesis is tenaciously discussed, additional evidence has yet to be amassed in support of this concept. It could, therefore, be possible that the RNA polymerase found in the cytoplasm in part represents a population of newly synthesized molecules. This and other hypotheses concerning the possible precursor relationships can be discussed faithfully only after additional experimental evidence has been obtained, since nothing is known at present about the mode of synthesis of this enzyme. ACKNOWLEDGMENTS We gratefully acknowledge financial support of the Deutsche Forschungsgemeinschaft and the able technical assistance of Misses D. Schwarz, and K. Eisenack as well as Mrs. Ch. Pfeiffer and F. Seifart in conducting the work outlined. Thanks are extended to Prof. Th. Wieland for having provided the c+amanitin and [%I-methyl-ramanitin, to Drs. Silvestri and Lancini (Gruppo Lepetit, Milano) for the rifamycin derivatives and to Dr. J. Chesterton for having donated a sample of myosin. In addition, we are thankful for the continued interest of Prof. P. Karlson to provide research facilities and to Prof. D. Gallwitz and C. E. Sekeris for numerous fruitful discussions. One of us (P. P. J.) is the recipient of a fellowship from the German Chemical Foundation. REFERENCES 1. (1970) Cold Spring Harbor symp. Quant. Biol. 66, 641-742. 2. ROEDER, R. G., AND RUTTER, W. J. (1969) Nature London 334, 234. 3. STIRPE, F., AND FIUME, L. (1967) Biochem.

J.

106, 779. 4. SEIFART, K. H., AND SEKERIS, C. E. (1969) 2. fvaturjorsch. 24b, 1638.

532

HEIFART,

BENECKE,

5. KEDINGER, C., GNUZDOWSKI, M., M~NDEL, J. L., GISSINGER, F., .IND CHAMBON, P. (1970) Biochem. Biophys. Res. Commun. 38, 165. 6. LINDELL, T., WEINBERG, F., MORRIS, P., ROEDER, R., .~ND RUTTER, W. (1970) Science 170,447. 7. JAKOB, S. T., SAJDEL, E. M., END MUNRO, M. N. (1970) Nature London 226, 60. 8. DEZELEE, S., SENTEN.~~, A., AND FROMAGEOT, P. (1970) Fed. Eur. Biochem. Sot. Lett. 7, 220. 9. NOVELLO, F., FIUMI, L., :~ND STIRPE, F. (1970) Biochem. J. 116, 177. 10. KEDINGER, C., NURET, P., AND CH~MBON, P. (1971) Fed. Eur. Biochem. Sot. Lett. 16, 169. 11. MANDEL, J. L., AND CHAMBON, P. (1971) Fed. Eur. Biochem. Sot. Lett. 16, 175. 12. WEAVER, R. F., BL.~TTI, S. P., AND RUTTER, W. J. (1971) Proc. Nat. Acad. Sci. U.S.A. 68, 2994. 13. CHESTERTOK, C. J., ,~ND BUTTERWORTH, P. (1971) Fed. Eur. Biochem. Sot. Lett. 12, 301. 14. SCHMUCKLER, E. A., .&ND TAT.~, J. R. (1971) Nature London 234, 39. 15. SAJDEL, E., AND JAKOB, S. T. (1971) Biochem. Biophys. Res. Commun. 46, 707. 16. SEIFART, K. H., :\ND SEKERIS, C. E. (1969) Eur. J. Biochem. 7, 408. 17. CH~CJVEAU, J., MOULE, Y., AND ROILLER, C. (1956) Exp. Cell Res. 11, 317. 18. BUSCH, H. (1967) in Methods in Enzymology (Grossmann, L., and Moldave, K., Eds.) Vol. 12, p. 448, Academic Press, New York. 19. WEBER, K., AND OSBORN, M. (1969) J. Biol. Chem. 244,4406. 20. BRAY, G. A. (1960) Anal. Biochem. 1, 279. 21. LUCK, J. M., RASMUSSEN, P. S., S.~T~KE, K., AND TSVETIKOV, A. N. (1958) J. Biol. Chem. 233, 1407. 22. HORGMN, P. A., AND GRIFFIN, D. H. (1971) Proc. Nat. Acad. Sci. U.S.A. 68,338. 23. SEIFART, K. H. (1970) Cold Spring Harbor Symp. Quant. Biol. 36, 719. 24. CHAMBON, P., GISSINGMR, F., MANDEL, J. L., KEDINGER, C., GNUZDOWSKI, M., AND MEIHLAC, M. (1970) Cold Spring Harbor Symp. Quant. Biol. 36, 693.

AND

JUHASZ

25. FURTH, J. J., HURWITZ, J., END ANDERS, M. (1962) J. Biol. Chem. 287. 2611. 26. EDMONDS, M., AND ABRAMS, R. (1962) J. Biol. Chem. 237, 2636. 27. GOTTESMAN, M. E., C~NNELAKIS, Z. N., .\ND C.4NNELBKIS, E. S. (1962) Biochim. Biophys. Acta 61, 34. 28. KLEMPERER, H. G. (1965) Biochim. Biophys. Acta 96, 251. 29. EDMONDS, M. (1965) J. Biol. Chem. 240, 4621. 30. HARDY, S., AND KURLAND, C. (1966) Biochemistry 6, 3668. 31. Twu, J. S., AND BRETTH:~UER, 12. K. (1971) Biochemistry 10, 1576. 32. HYATT, E. A. (1967) Biochim. Biophys. Acta 142, 246. 33. G.~D.~LETA, M., GRECO, M., AND SACCONE, C. (1970) Fed. Eur. Biochem. Sot. Lett. 10, 54. 34. REID, B. D., AND PARSONS, P. (1971) Proc. Nat. Acad. Sci. U.S.A. 68, 2830. 35. K~NTZEL, H., AND SCHAFER, K. P. (1971) AVature London AVew Biol. 231,265. 36. WINTNRSBERGER, E., AND WINTERSBERGER, U. (1970) Fed. Eur. Biochem. Sot. Lett. 6, 58. 37. KILT, G. E., AND FAUST, A. S. (1969) Arch. Biochem. Biophys. 134, 103. 38. CHESTERTON, C. J., HUMPHREY, S., .\ND BUTTERWORTH, P. (1972) Biochem. J. in press. 39. SIEBERT, G., personal communication. 40. STEIN, H., AND HAUSEN, P. (1971) Eur. J. Biochem. 14, 270. 41. SCHLAB~CH, A., FRIDLENDER, B., BOLDEN, A., AND WEISSBACH, A. (1971) Biochem. Biophys. Res. Commun. 44, 879. 42. ROYCHOUDHURY, R., AND BLOCH, I>. (1969) J. Biol. Chem. 244, 3359. 43. CHANG, L. M., AND BOLLUM, F. J. (1971) J. Biol. Chem. 246, 5835. 44. HAINES, M. E., JOHNSTON, I. R., END MATHIAS, A. P. (1970) Fed. Eur. Biochem. Sot. Lett. 10, 113. 45. WEISSBACH, A., SCHLABACH, A., FKIDLENDER, B., .\ND BOLDEN, A. (1971) Nature London New Biol. 231, 167. 46. BELL, E. (1971) Science 174, 603. 47. GALLXVITZ, I)., .\ND MUELLER, G. (1969) Eur. J. Biochem. 9, 431.