DNA-dependent RNA-polymerases from Artemia embryos

DEVELOPMENTAL

BIOLOGY

45, 34-43 (19%)

DNA-Dependent Characterization HOWARD Department

C.

from Artemia

RNA-Polymerases

of Polymerases I and II from Nauplius Larvae

BIRNDORF,

of Biochemistry,

Embryos

JAMES D’ALESSIO,

Wayne State University Accepted

February

AND JOSEPH

School

C.

BAGSHAW’

of Medicine Detroit, Michigan

48201

14, 1975

Partial purification and characterization of DNA-dependent RNA-polymerases from nauplius larvae of the brine shrimp, Artemia salina, are described. Fractionation of solubilized RNApolymerases on columns of DEAE-cellulose yielded partially purified preparations of RNA polymerases I and II. The properties of these enzymes were found to be similar to properties of corresponding enzymes from other animal sources. A significant change in the relative amounts of polymerases I and II occurs between 36 and 72 hr of development. Polymerase activity obtained from 36-hr nauplii consisted of approximately equal amounts of polymerases I and II, whereas polymerase II accounted for more than 80% of the activity recovered from 72-hr nauplii. Total polymerase activity was lower at 72 than at 36 hr. The significance of these changes in relation to the decrease in RNA synthesis in uiuo that occurs after 36 hr is discussed. INTRODUCTION

the distribution of multiple RNA polymerases was essentially constant despite major changes in the pattern of transcription. Even among simpler developmental systems conflicting results have been found. For example, Sol1 and Fulton (1974), studying the ameboflagellate Naegleria, found no change in the amount or distribution of polymerases during a period of alteration in the pattern of RNA synthesis. In contrast to this, Pong and Loomis (1972) detected a 75% decrease in the amount of RNA polymerase II per cell in differentiating Dictyostelium but found no difference in the relative proportions or properties of multiple polymerases. From these studies it is apparent that alterations in the pattern of RNA synthesis in uiuo are not always accompanied by changes in RNA polymerases that are amenable to in vitro experimentation. Each biological system of interest to developmental biologists must be examined in this respect. The brine shrimp, Artemia salina, is an excellent biological system for biochemical studies of differentiation. The dormant, encysted gastrulae (cysts) are available at any time in large quantity. In artificial sea

Embryonic development is characterized by closely regulated changes in patterns of gene expression. Some of these changes occur at least in part at the level of transcription. Since the discovery that eukaryotes possess multiple forms of DNAdependent RNA-polymerase (Roeder and Rutter, 1969, 1970a), considerable attention has been given to the possibility that changes in the transcription pattern are correlated with changes in the amounts and/or distribution of multiple RNA polymerases. In some cases such correlations have been found. For example, Roeder and Rutter (1970b) demonstrated that alterations in RNA synthesis in developing sea urchin embryos were accompanied by changes in the relative proportions of the RNA polymerases. The observed differences in the polymerases agreed with the putative function of these enzymes in synthesis of ribsomal and nonribosomal RNA (Roeder and Rutter, 1970a). On the other hand, Roeder et al. (1970; Roeder, 1974b) found that in developing Xenopus embryos ‘To whom dressed.

requests

for reprints

should

be ad34

Copyright 0 1975 by Academic Press, Inc. All rights of reproduction in any form reserved.

BIRNDORF,

D’ALESSO AND BAGSHAW

water the embryos will develop into nauplius larvae in a fairly synchronous manner. During this period of postgastrula development, changes in the metabolism of nucleotides (Finamore and Clegg, 1969) and nucleic acids (McClean and Warner, 1971) take place. McClean and Warner (1971) found that maximal incorporation of nucleotides into RNA occurred in freeswimming nauplius larvae 30-36 hr after encysted gastrulae resumed development. Thereafter the rate of nucleotide incorporation decreased to less than 10% of the maximal rate. These authors further showed that uptake of radioactive precursors into intracellular nucleotide pools did not diminish over this period. Thus it appears that Artemia larvae undergo a dramatic reduction in the rate of RNA synthesis. We have initiated studies directed at understanding the molecular basis of this major quantitative change in transcription, and as a first step we have isolated DNA-dependent RNA-polymerases I and II from nauplii at stages before and after the reduction in RNA synthesis. Each form of the enzyme has distinct requirements for maximal acitivity, and these requirements are independent of the age of the nauplii. A marked alteration in relative amounts of the two forms occurs, polymerase I being the major component at 36 hr while polymerase II predominates at 72 hr. MATERIALS

AND

METHODS

Preparation of Artemia nauplii. Encysted gastrulae (Salt Lake variety, Long Life Fish Food Company) were sterilized by treatment with 7% antiformin solution (Nakanishi et al., 1962). This treatment removed cryptobiotic organisms which contaminate commercial preparations of cysts. The washed cysts (10 g, wet wt) were then added to 2-l Erlenmeyer flasks with 500 ml of sterile sea water (McClean and Warner, 1971) containing penicillin G and streptomycin sulfate at 1,000 units/ml and 1 mg/ml, respectively. Flasks were incu-

RNA Polymerase

from Artemia

Embryos

35

bated at room temperature with constant gentle shaking to ensure adequate oxygenation. After the desired period of incubation, the free-swimming nauplius larvae were harvested by emptying the contents of the flask into a l-l separatory funnel, the upper part of which was covered so that the positively phototropic nauplii would gather at the stopcock. The larvae were then gently drained onto Whatman #l filter paper and washed with distilled water. All subsequent procedures were performed at 4°C unless specified otherwise.

Isolation of nuclei and solubilization of RNA polymerase. The nauplii were suspended in large Dounce homogenizers containing ice-cold homogenizing medium consisting of 0.01 M Tricine’ (pH 8.0), 5 mM EDTA, and 0.25 M sucrose (about 0.5 g of nauplii/ml of medium). The larvae were homogenized with ten strokes each of the loose and tight pestles. The homogenate was filtered through glass wool to remove shell fragments and centrifuged at l,OOOg for 10 min. The resulting pellet was resuspended in homogenizing medium by ten strokes of the tight pestle and the suspension was centrifuged at 750g for 10 min. RNA polymerase was solubilized from the resulting pellet by a modification of the procedure of Roeder and Rutter (1970a). The pellet was resuspended in homogenizing medium (typically, in 1 ml for the pellet from 2 culture flasks) and made 0.3 M in (NH,),SO, by addition of a suitable volume of 4 M (NH,),SO,, pH 8.0. The resulting viscous suspension was sonicated for 25 set in 5-set intervals using the large probe of a Biosonic sonicator operated at full power. The sonicate was diluted with two volumes of TMD buffer (20 mM Tritine, pH 8.0, 1 mM dithiothreitol, 5 mM MgCl,, 0.5 mM EDTA, 20% glycerol, by volume) and centrifuged for 20 min at 30,OOOg.The nucleoprotein pellet was discarded and the pale orange supernatant fraction was diluted with one volume of ‘Tricine

is N-tris(hydroxymethyl)methyl

glycine.

36

DEVELOPMENTALBIOLOGY

vOLUME45,1975

TMD buffer to bring the final (NH,),SO, DNA preparations. DNA was extracted concentration to 0.05 M. from encysted embryos by the method of DEAE-cellulose chromatography. RNA Kay et al. (1952) and further purified by polymerase was fractionated by chromaPronase treatment and extraction with tography on columns of DEAE-cellulose chloroform-isoamyl alcohol, as described immediately after solubilization. For the by Schildkraut and Maio (1969). Calf thyamount of nauplii obtained from two cul- mus DNA was purchased from Sigma ture flasks, a 5-ml bed volume was suffiChemical Company. Bacillus subtilis DNA cient for proper fractionation of the enzyme was a generous gift of Dr. Veronica Maher activity. The DEAE-cellulose (Whatman of the Michigan Cancer Foundation. DNA DE52) was suspended in TMD buffer con- preparations were denatured by treatment taining 0.04 M (NH,),SO, (start buffer) with NaOH at pH 12. and equilibrated with two bed volumes of the same buffer. The soluble enzyme prepRESULTS aration was applied to the surface of the Chromatographic Fractionation of Solubicolumn bed and fraction collection was lized RNA Polymerases begun immediately. After all the enzyme RNA polymerases solubilized from crude had been applied, the column was washed with one bed volume of start buffer and nuclear preparations obtained from nauplii at the peak of RNA synthetic activity (36 then developed with a 50-ml linear gradihr) were fractionated on columns of DEAEent of 0.04-0.5 M (NH,),SO, in TMD cellulose. A typical fractionation is illusbuffer at a flow rate of 10 ml/hr. Forty trated in Fig. 1. The major portion of fractions were collected and assayed immediately for RNA-polymerase activity. Rela- protein failed to adhere to the column, while RNA-polymerase activity was adtive protein concentrations of the fractions and then eluted with were estimated by measuring the absorb- sorbed quantitatively a salt gradient to yield two peaks. The first ance at 280 nm in a Beckman grating peak of activity was eluted at 0.10-0.15 M spectrophotometer. Fractions containing and the second at 0.20-0.25 M RNA polymerases I and II were pooled (NH,),SO, When coiumn fractions were separately, mixed with an equal volume of (NH,),SO,. assayed in the presence of the mushroom glycerol, and stored at -80°C. toxin a-amanitin (2 pg/ml), the first peak Assay for RNA-polymerase activity. RNA-polymerase activity was assayed by was resistant and the second was commeasuring incorporation of [3H JUTP into pletely inhibited. Thus the first and second peaks of activity correspond to RNA poRNA as described elsewhere (Bagshaw, lymerase I and II, respectively, as isolated 1974). Assay mixtures (125 ~1) contained the following: 50 mM Tricine, pH 8.0, 5 from other eukaryotic sources. No signifiwas found in mM dithiothreitol, 5 mM creatine phos- cant polymerase activity flow-through fractions or in fractions elutphate, 50 pglml of creatine phosphokinase, ing after polymerase II. 0.5 mM each ATP, CTP, GTP, 0.05 mM [SH]UTP (0.5 mCi/mmole), and 25 kg of Ionic Requirements of RNA Polymerases I denatured calf thymus DNA. In addition, and II assays of column fractions contained 2 mM MnCl, and 25 cl1 of the column fraction. Chromatographically separated RNA Concentrations of divalent cations and polymerases I and II were assayed to deter(NH,),SO, in other assays are given in the mine the ionic conditions required for maxappropriate figures and tables. Assays imal activity of the two enzymes. The were incubated for 30 min at 27°C. optimal concentration of (NH,) &SO, was

BIRNDORF,

D’ALEWO

RNA Polymerase

AND BACSHAW

from Artemia

Embryos

37

IO

6

IO

20 FRACTION

30

40

NUMBER

FIG. 1. Fractionation of RNA polymerase from 36-hr nauplii on DEAE-cellulose. 0, RNA-polymerase activity; A, polymerase activity in the presence of a-amanitin (2 pplml); - - -, absorbance at 280 nm; --, estimated (NH,),SO, concentration.

found to be 30-40 mM for polymerase I and loo-120 mM for polymerase II (Fig. 2). Both polymerases required a divalent cation for activity, and both were more active with Mn2+ than with Mg2+ (Fig. 3). The optimal concentration of Mn2+ was determined to be 2 mM. Addition of 10 mM Mg2+ in the presence of 2 mM Mn2+ inhibited RNA polymerase I by 20% and RNA polymerase II by 5% (data not shown). Polymerase activities varied only slightly between pH 7.0 and 9.0, with maximal activity at about pH 8.0 (Fig. 4). Comparison plates

of Different

DNA’s

as Tem-

Native (double-stranded) and denatured (single-stranded) forms of DNA from Artemia cysts, calf thymus, and Bacillus subtilis were tested as templates for RNA polymerases I and II from 36-hr nauplii.

Results are presented in Table 1. Denatured forms of all three DNA’s were better templates than the corresponding native forms, and, with either Artemia or calf thymus DNA, polymerase II showed a more marked preference for denatured DNA than did polymerase I. Artemia DNA was a slightly better template than calf thymus DNA, except in the case of polymerase II with native DNA’s. Native B. subtilis DNA was a very poor template for polymerase I and was substantially poorer than eukaryotic DNA’s for polymerase II. However, the denatured form of B. subtilis DNA was as good a template as calf thymus DNA for either enzyme. Effect of lnhibitors Polymerases

on Separated

RNA

a-Amanitin, cycloheximide, and rifampicin were tested for the ability to inhibit

DEVELOPMENTALBIOLOGY

I 40

a0 (NH4J22.04

120

160

200

(“IM)

FIG. 2. Activity of RNA polymerases I and II as a function of (NH,)SO, concentration. All assay solutions contained 2 mM MnCl,. Peak fractions of polymerase I and II were diluted and assayed at the indicated salt concentration. A, RNA polymerase I; 0, RNA polymerase II.

vOLUME45,1975

this decrease, polymerase from 36 and 72hr nauplii were fractionated simultaneously on twin DEAE-cellulose columns (Bagshaw, 1974). The results of such a fractionation are shown in Fig. 5. At 72 hr, the peak of polymerase II activity was substantially greater than the peak of polymerase I, while at 36 hr the polymerase I activity predominated (cf. Fig. 1). In this experiment, polymerases were extracted from the same number of cultures at 36 and 72 hr. Thus the total activity recovered at 72 hr was significantly less than the activity recovered at 36 hr. No difference in the 4

A

Artemia RNA-polymerase partially purified by DEAE-cellulose chromatography (Table 2). Inhibition of polymerase II by a-amanitin was detected at concentrations as low as 0.01 pg/ml, and was essentially complete at 0.1 pug/ml. Polymerase I was resistant to cy-amanitin at all concentrations tested. Cycloheximide, which has been reported to inhibit RNA polymerase I from some sources (Horgen and Griffin, 1971; Timberlake et al., 1972a,b; Weissman, 1973), had very little inhibitory effect upon Artemia polymerase I and none upon polymerase II. Neither polymerase from Artemia was inhibited by rifampicin, which is selective for prokaryotic RNApolymerases and possibly for mitochondrial RNA-polymerases (Gadeleta et al., 1970; Kuntzel and Schafer, 1971; Reid and Parsons, 197 1). Comparison of RNA Polymerases Nauplii of Different Ages

from

The decrease in RNA synthesis in viva reported by McClean and Warner (1971) is substantially complete by 72 hr after immersion of cysts in sea water. In order to compare the relative proportions of polymerases I and II at stages before and after

4

8 DIVALENT

12 CATION

16

20

lmM)

FIG. 3. Activity of RNA polymerases I and II as a function of divalent cation concentration. Peak fractions of polymerase I (A) and polymerase II (B) were assayed at 30 and 120 mM (NH,),SO,, respectively, in the presence of the indicated concentrations of MgCl, (0) or MnCl, (0).

BIRNDORF, D’ALESSIO AND BACSHAW

39

RNA Polymerase from Artemia Embryos

estimated that the true activity of polymerase II was about twice what it appears to be in Fig. 1. Thus, 36-hr nauplii yielded roughly equal amounts of polymerases I and II. RNA polymerases I and II were by far the major components found in Artemia preparations. A small shoulder of amanitinresistant activity which was frequently seen eluting between polymerases I and II may represent a subfractionation of polymerase I. No activity comparable to polym70

SO

9.0

TABLE

PH

FIG. 4. Activity of RNA polymerases I and II as a function of pH. Peak fractions of polymerase I (01 and polymerase II (0) were assayed in the presence of 30 and 120 mM(NH,),SO,, respectively, and 2 mM MnCl,.

elution position of either enzyme was observed. DICUSSION

Developing nauplius larvae of Artemia, like other eukaryotic systems, contain at least two forms of DNA-dependent RNApolymerase. It should be pointed out that the preparations of nuclei used in this study contained large amounts of yolk platelets, due to the difficulty in separating nuclei and yolk platelets by aqueous methods. Nevertheless, good yields of soluble RNA-polymerases were obtained. Fractionation of polymerases on DEAEcellulose revealed two major peaks of activity that correspond to RNA polymerases I and II (Fig. 1). The activity of polymerase I appears to be much greater than that of polymerase II, but this is not actually the case. Due to the salt gradient used to develop the column, fractions containing polymerase I were assayed at concentrations of (NH,),SO, that were approximately optimal for that form, whereas fractions containing polymerase II were assayed at decidely suboptimal concentrations. From the data in Fig. 2 it can be

1

COMPARISONOF DIFFERENT DNA’s AS TEMPLATE FOR Artemia RNA-POLYMERASES” UMP incorporated (cpm)

DNA used as template

Artemia, native Artemia, denatured Calf thymus, native Calf thymus, denatured Bacillus subtilis, native Bacillus subtilis, denatured

Polymerase I

Polymerase II

7,150 11,475 6,004 9,314 499 9,257

2,612 889 2,039 300 2,032

873

“All assays contained 2 mM MnCl, and 10 c(g of the indicated DNA. Polymerases were prepared from 36 hr nauplii. Assays contained (NH,),SO, at either 30 mM (polymerase I) or 120 mM (polymerase II). TABLE

2

EFFECT OF INHIBITORS ON Artemia RNA-POLYMERASEV Inhibitor

Percent inhibitionb

I

Polymerase II

0 1 1 1 5 0

56 84 93 98 1 0

Polymerase 1

a-Amanitin, 0.01 rig/ml a-Amanitin, 0.05 fig/ml a-Amanitin, 0.10 rig/ml cu-Amanitin, 1.0 @g/ml Cycloheximide, 500 &ml Rifampicin, 100 &ml

“Assay mixtures contained 25 pg of denatured calf thymus DNA, 2 mM MnCl,, and (NH,),SO, at 30 mM (polymerase I) or 120 mM (polymerase II). b Control activities were 3,466 cpm for polymerase I and 5,406 cpm for polymerase II.

40

DEVELOPMENTALBIOLOGY

VOLUME 45,1975

-2 .O

II.5 - I .O

.s

: 2

-2 1.0

- I .s

- I .O

I

IO

20 F RAGTION

30

.s

40

NUMBER

FIG. 5. Comparison of RNA polymerase from nauplii at different developmental stages. Preparations obtained from nauplii at 72 hr (A) and at 36 hr (B) were fractionated simultaneously on twin DEAE-cellulose columns. 0, RNA-polymerase activity; - - -, absorbance at 280 nm; --, estimated (NH,),SO, concentration.

erase III has ever been observed. However, in view of the notorious instability of this enzyme, and the recent reports that it is much more active with poly[d(A-T) ] as a template (Roeder, 1974a), it is impossible to rule out the existence of polymerase III in Artemia. Moreover, Sergeant and Krsmanovic (1973) found that RNA polymerase III could be detected in extracts of KB cells after fractionation of DEAESephadex but not on DEAE-cellulose. Further purification of Artemia RNA-polymerases may reveal heterogeneity not appar-

ent at this stage, possibly including the presence of RNA polymerase III. The conditions required for maximal activity of Artemia RNA-polymerases I and II (Figs. 2-4, Table 1) are generally similar to optimal conditions reported for the corresponding enzymes from other sources (reviewed by Jacob, 1973). However, certain minor differences should be noted. RNA polymerase I from most sources prefers Mg*+ as the divalent cation and native DNA as the template. In contrast to this, both Artemia polymerases I

BIRNDORF,

D’ALESSIO

AND BACSHAW

and II were more active with Mn2+ and denatured DNA. Substituting Mg2+ for Mn2+ in the assays did not alter the preference of either polymerase for denatured DNA (unpublished observation). The slight inhibition of the polymerases by Mg2+ in the presence of Mn2+ probably represents simple competition between the ions, the enzymes being slightly less active when the less favored ion is bound. The properties of Artemia polymerase II with respect to ionic optima and template preference are quite similar to the properties of virtually every polymerase II reported to date. When tested with DNA’s from different sources, RNA polymerases I and II showed only a slight preference for Artemia DNA as template in comparison with calf thymus DNA. Bacillus subtilis DNA, however, was a very poor template in the native form, but was as good as calf thymus DNA when the bacterial DNA was in the denatured form. A speculative, but plausible, explanation for the template activity of B subtilis DNA is that in its native state it contains relatively few sites at which the eukaryotic polymerases can bind and/or initiate transcription. Thus the native DNA would be an inefficient template. The denatured form, on the other hand, is as good a template as eukaryotic DNA’s because enzyme binding and initiation presumably occur randomly on singlestranded DNA, As anticipated from studies in other eukaryotic systems, Artemia RNA-polymerase I was resistant to the mushroom toxin a-amanitin at 2 pg/ml, whereas polymerase II was completely inhibited at this concentration (Fig. 1). It is interesting to compare the effect of lower concentrations of a-amanitin on Artemia polymerase II (Table 2) with results reported for this enzyme from other animals, particularly arthropods. Doenecke et al. (1972) found that inhibition of RNA polymerase II from the blowfly, Calliphora erythrocephala, re-

RNA Polymerase

from Artemia

Embryos

41

quired a tenfold higher concentration of a-amanitin than that needed to achieve equal inhibition of rat-liver polymerase II. In contrast to this, polymerase II from Drosophila melanogaster embryos has been reported to be as sensitive to a-amanitin as mammalian polymerase II (Phillips and Forrest, 1973). In this respect Artemia RNA-polymerase II more closely resembles the Drosophila and mammalian enzymes. Inhibition of Artemia polymerase II is nearly complete at 0.1 pug/ml of LYamanitin, a concentration at which the analogous enzyme from Calliphora is inhibited by about 60% (Doenecke et al., 1972). The effects of cycloheximide and rifampicin on Artemia RNA-polymerase I were also tested (Table 2). Cycloheximide, which has been reported to inhibit RNA polymerase I from some protists (Horgen and Griffin, 1971; Timberlake et al., 1972a,b; Rudick and Weissman, 1973), had very little effect upon Artemia polymerase I and none upon polymerase II. Both enzymes were also resistant to rifampicin, an inhibitor of prokayotic polymerases, and in some cases also reported to be an inhibitor of mitochondrial RNA-polymerases (Gadeleta et al., 1970; Kuntzel and Schafer, 1971; Reid and Parsons, 1971). Thus the Artemia polymerase preparations used in this study were not contaminated by detectable amounts of bacterial RNA polymerases. However, mitochondiral contamination cannot be clearly ruled out due to conflicting reports regarding the sensitivity of mitochondrial RNA polymerases to rifampicin. Two changes occur in Artemia RNA polymerases between 36 and 72 hr after immersion of the cysts. As shown in Fig. 5, the total amount of polymerase activity recovered from 72.hr nauplii is roughly one-tenth (polymerase I) to one-half (polymerase II) the activity from 36 hr (note the difference in scale in the figure). This apparent loss of polymerase activity is not

42

DEVELOPMENTALBIOLOGY

surprising in view of the reduction in RNA synthesis in uiuo that occurs at this time (McClean and Warner, 1971), although RNA-polymerase levels do not necessarily reflect rates of RNA synthesis. A more striking difference is the change in the relative amounts of polymerase I and II. As mentioned previously, the actual activity of polymerase II assayed under optimal conditions is about twice what it appears to be in the column fractions. Thus RNApolymerase activity recovered from 72-hr nauplii comprises more than 80% polymerase II, whereas polymerase activity from 36-hr nauplii is approximately equally distributed between I and II. The differences in relative activity might be due to the presence of factors other than RNA polymerases in these preparations; this seems unlikely, however, because chromatography of mixed preparations of polymerases from 36 and 72-hr nauplii yielded polymerases I and II in the amounts predicted by simple mixing (unpublished observations). The biological significance of this shift in relative amounts of polymerases I and II is not clear, since McClean and Warner have reported that synthesis of all classes of RNA diminishes from 36 to 72 hr. However, it should be pointed out that preparation of RNA polymerases from subnuclear fractions is not currently feasible with Artemia. Therefore it can only be assumed that polymerase I is the nucleolar form and polymerase II the nucleoplasmic. In summary, the studies presented here indicate that the brine shrimp Artemia salina is a useful biological system for investigating the function of RNA polymerases in development. Polymerases I and II have been isolated from Artemia nauplius larvae and shown to have properties similar to corresponding enzymes from other eukaryotes. The reduction in RNA synthesis in viuo that occurs in developing nauplii is accompanied by a reduction in total RNA-polymerase activity as well as a major change in the relative amounts of

VOLUME 45,1975

polymerases I and II. Experiments are now in progress to define more precisely the time at which this change takes place and to determine whether or not it involves an alteration in the subunit structure or other physical parameters of these enzymes. This work was supported by Research Grant No. GM21376-01 from the National Institute of General Medical Sciences and by a Faculty Grant-in-Aid from Wayne State University. The authors wish to thank Mrs Bemice Bond for her excellent technical assistance. REFERENCES BAGSHAW, J. C. (1974). Biochem. Biophys. Res. Commun. 57, 177-182. DOENECKE, D., PFEIFFER, C., and SEKERIS, C. E. (1972). FEBS Lett. 21, 237-243. FINAMORE, F. J., and CLEGG,J. S. (1969). In “The Cell

Cycle: Gene-Enzyme Interactions” (G. M. Padilla G. L. Whitson, and I. L. Cameron, eds.), pp. 249278 Academic Press, New York. GADELETA, M. N., GRECO, M., and SACCONE,C. (1970). FEBS Lett. 10, 54-56. HORGEN, P. A., and GRIFFIN, D. H. (1971). Proc. Nut. Acad. Sci. USA 68, 338-341. KAY, E. R. M., SIMMONS, N. S., and DOUNCE, A. L. (1952). J. Amer. Chem. Sot. 70, 1724-1726. KUNTZEL, H., and SCHAFER, K. (1971). Nature New Biol. 231, 265-269. MCCLEAN, D. K., and WARNER, A. H. (1971). Deuelop. Biol. 24, 88-105. NAKANISHI, Y. H., IWASAKI, T., OKIGAKI, T., and KATO, H. (1962). Annot. Zool. Jap. 35, 223-228. PHILLIPS, J. P., and FORREST, H. S. (1973). J. Biol. Chem. 248, 265-269. PONG, S. S., and LOOMIS, W. F. (1973). J. Biol. Chem.

248, 3933-3939. REID, B. D., and PARSONS,P. (1971). Proc. Nat. Acad. Sci. USA 68, 2830-2834. ROEDER, R. G. (1974a). J. Biol. Chem. 249, 241-248. ROEDER, R. G. (197413).J. Biol. Chem. 249, 249-256. ROEDER, R. G., REEDER, R. H., and BROWN, D. D. (1970). Cold Spring Harbor Symp. Quant. Biol. 35,

725-735. ROEDER, R. G., and RUTTER, W. J. (1969). Nature (London) 224, 234-237. ROEDER, R. G., and RUT~ER,W. J. (1970a). Proc. Nat. Acad. Sci. USA 65, 675-682. ROEDER, R. G., and RUTTER, W. J. (1970b). Biochemistry 9, 2543-2553. RUDICK, V. L., and WEISSMAN, R. A. (1973). Biochim. Biophys. Acta 299, 91-102. SCHILDKRAUT, C. L., and MAIO, J. J. (1969). J. Mol. Biol. 46, 305-312.

BIRNDORF, D’ALESSIO AND BAGSHAW SERGEANT,A., and KRSMANOVIC, V. (1973). FEES Lett. 35, 331-335. SOLL, D. R., and FULTON, C. (1974). Develop. Biol. 36, 326-244. TIMBERLAKE, W. E., HAGEN, G., and GRIFFIN, D. H.

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(1972). Biochem. Biophys. Res. Commun. 48, 823-827. TIMBERLAKE, W. E., MCDOWELL, L., and GRIFFIN, D. H. (1972). Biochem. Biophys. Res. Commun. 46, 942-947.