Events surrounding the early development of Euglena chloroplasts

ARCHIVES

OF

Events

BIOCHEMISTRY

AND

Surrounding

Photoregulation

BIOPHYSICS

for Photobiology

201-216

(1976)

the Early Development of the Transcription Ribosomal

DAN Institute

177,

of Cells

COHEN3 and

Organelles, Received

of Chloroplastic RNAs’, 2

JEROME

AND

of Euglena

Brandeis January

Chloroplasts

and Cytoplasmic

A. SCHIFF4 University,

Waltham,

Massachusetts

02154

28, 1976

During light-induced chloroplast development in dark-grown resting cells of wild-type Euglena gracilis Klebs var. bacillaris Pringsheim, plastid ribosomal RNA (pl-rRNA) increases from about 2 to 25% of the total cellular RNA; concomitantly, 32POqs~ is incorporated linearly from 0 to 48 h. The net amount of cytoplasmic ribosomal RNA (cytrRNA) does not change during light-induced chloroplast development in wild type, but .12P0,3m is incorporated with a peak rate at 24-36 h, indicating that turnover of cyt-rRNA takes place during light-induced plastid development. The extent of this turnover is about 60% by 36 h of development. Such light-induced turnover of cyt-rRNA also occurs in resting cells of dark-grown W,BUL, a mutant of Euglena which lacks detectable plastid DNA and protochlorophyll(ide), with similar kinetics but to one-third the extent of wild-type cells. Labeling of cyt-rRNA at 3 h of development is primarily in the polysome fraction compared with the monosomes, indicating that newly synthesized ribosomes are immediately active in protein synthesis. Labeling of pl-rRNA continues at the same rate in darkness after a brief light induction while cyt-rRNA labeling decreases 50% every 3 h in darkness after a brief light induction. Labeling of cyt-rRNA in wild type and W,BUL is induced most effectively by blue light, green and red light being relatively ineffective. Labeling of pl-rRNA in wild-type cells is induced most effectively by blue light and red light, and the ratio of effectiveness of blue to red to green is the same as the ratios of absorption in these regions of the protochlorophylhide) holochrome from beans. These results are consistent with other data which indicate that the nonplastid aspects of plastid development are under control of a nonchloroplast blue light receptor while the plastid events are under the control of the plastid-localized protochlorophyll(ide) system.

The developing chloroplast of Euglena var. bacillaris requires contributions of energy, reducing power, small

molecules, and large molecules from the rest of the cell (18). While certain constituents arise from synthetic activities within the developing chloroplast, photosynthesis is not required for plastid development and several plastid proteins are known which are coded outside the plastid DNA (probably in nuclear DNA) and are synthesized outside the chloroplast. From this and other information, it appears that the lag period which occurs during the first 12 h of

gracilis

’ Supported by Grant GM 14595 from the National Institutes of Health. This is paper No. 10 in a series, Paper No. 9 appeared in Planta 131:1-g. * Work similar to that presented in this paper has been reported recently by Dr. Martin Steup, Pflanzenphysiol. Institut d. Univ. Goettingen, Germany, in Plant Physiology Vol. 57, Number 5, (1976) May, entitled, “Light dependent regulation of ribosomal and transfer RNA synthesis”. s Goldwyn Fellow of the University. This work was taken from a thesis submitted to the Graduate Faculty of Brandeis University in partial fulfillment of the requirements for the Ph.D. Degree. Present

address: Department of Applied Genetics, University, Jerusalem, Israel. 4 Abraham and Etta Goodman Professor ogy; author to whom reprint requests should 201

Copyright All rights

0 1976 by Academic Press, of reproduction in any form

Inc. reserved.

Hebrew of Biolbe sent.

202

COHEN

AND

plastid development in dark-grown resting cells exposed to light is a time in which the plastid receives a major contribution from the cytoplasm; beyond 12 h the major synthetic activities appear to take place within the plastid itself. The two aspects of plastid development comprising plastid and nonplastid events are controlled by at least two different light receptors, a receptor absorbing mainly in blue light which controls nonplastid events and a plastidlocalized protochlorophyll(ide) system absorbing in both the blue and red regions of the spectrum which controls chlorophyll synthesis and other plastid-localized events (18, 27, 30, 31, 35). Since the light-induced development of the proplastid into the chloroplast in darkgrown resting cells of Euglena requires extensive synthesis of plastid proteins which takes place in both the cytoplasm and chloroplast using information contained in nuclear and plastid DNAs (18, 20), it is of interest to inquire into the formation of the protein-synthesizing machinery of these two compartments during the developmental process. In this paper we report the results of experiments in which the synthesis of the ribosomal RNAs of the chloroplast and cytoplasm are measured during chloroplast development. We show that the transcription of cytoplasmic and plastidic ribosomal RNAs are under control by separate pigment systems and provide additional information concerning the regulation of rRNA formation in wild-type Euglena cells and in a mutant in which plastid DNA is undetectable. A brief abstract of this work has already appeared (1). MATERIALS

AND

METHODS

Culture conditions. Euglena gracilis Klebs var. bacillaris Pringsheim, maintained either in the light or in darkness for many years, was grown on Hutner’s pH 3.5 medium (2). All manipulations of dark-grown cells were done under green safelights (3). Since many of the experiments to be described involved the use of radioactive phosphate, a resting medium was devised containing a reduced level of nonradioactive phosphate and a pH of 4.5 which was found to be optimal for phosphate uptake. This medium allowed normal light-induced chloroplast development in nondividing cells. The modified resting medium contained: mannitol (0.05 M), MgCl,

SCHIFF (0.01 M), KH,PO, (0.00013 M) and 1,2,3,4-cyclopentane-tetracarboxylic acid (“cycle acid” (Aldrich)) (0.01 M). Cycle acid buffer (48) allowed the use of more concentrated cell suspensions than usual, and normal chlorophyll formation was obtained at cell densities up to 4 x 10’Vml. To avoid carryover of phosphate from the growing medium, the method of preparing resting cells was also modified to include a centrifugation step. All steps were carried out aseptically. Two hundred milliters of a growing culture containing 5-7 X lo5 cells/ ml were centrifuged at room temperature for 3 min at 800g. The growth medium was discarded and the pellet was resuspended in sterile cycle acid resting medium at the same cell concentration as in the growth medium. The cells were allowed to cease division in the dark before exposure to light for chloroplast development. The centrifugation step was found to have little or no effect on subsequent light-induced plastid development. Lighting conditions. Chloroplast development was carried out at 150 fc of light provided by G.E. daylight and red fluorescent tubes (#7212r) (5). This light source provided up to 600-fc intensity for certain experiments, and lower intensities were obtained by using layers of gauze as neutral density filters. For experiments using selected regions of the visible spectrum, light intensities were measured with a Weston Sunlight Illumination meter calibrated against a thermopile (Eppley) in each spectral region. Red light (610-720 nm) was provided by a G.E. fluorescent tube (#7212r) masked at the ends with black electrician’s tape where the phosphor becomes unreliable. Blue light (400-510 nm) was provided by daylight white fluorescent tubes filtered by blue Plexiglas (Roehm and Haas #2424). Green light (520-570 nm) was provided by daylight fluorescent tubes filtered by Wratten filters #16 and #61. Conditions for Labeling with 32P0,3-. Sterile carrier-free H,32P0,3(32P,)” was added directly to the experimental cultures to a final level of 0.1 mCi/ml yielding a final specific activity of 0.08 mCi/pmol of phosphate. Since no lag in uptake was found, the isotope was added at time zero of the experiments. Since phosphate uptake greatly exceeded labeling of RNA, the extent of RNA labeling was not limited by phosphate uptake, and ample phosphate was present in the medium to provide a nonlimiting reservoir throughout the experiments. That RNA was uniformly labeled under these conditions is indicated by the finding that in experiments where degradation of RNA was allowed to occur through omis5 Abbreviations used: V 32P0.,3m; cyt-rRNA, cytoplasmic (nuclear) ribosomal RNA; pl-rRNA, chloroplast or proplastid ribosomal RNA; sp act, specific activity (counts per minute per microgram of RNA); Pi-N pool, phosphate and nucleotide pool; SDS, sodium dodecyl sulfate; DNase, deoxyribonuclease.

PHOTOREGULATION

OF

sion of salt or thioglycollate, the 10 or more degradation fractions obtained had fairly uniform specific activities compared with each other. RNA extraction. The procedure used was a combination of the techniques employed by Rawson and Stutz (6) and Heizmann (7). All manipulations were carried out at 4°C. Twenty-five milliliters of resting culture was transferred to a 30-ml Corex glass tube, which was centrifuged at 8OO.g for 5 min in a refrigerated Sorvall centrifuge (RCZ-B). The medium was discarded and the cells were washed once with 10 ml of buffer I (100 ml of Tris-HCl (0.01 M), pH 7.6, MgCl, (0.002 M), KC1 (0.01 M)) and centrifuged again at 800g for 5 min. The cell pellet was resuspended in 3 ml of buffer I supplemented with 20 mg/ml of sodium thioglycollate and was agitated well on a Vortex mixer and was then allowed to stand for 2 min in an ice bucket. Lysis was effected by adding 0.2 ml of 20% w/v in water recrystallized sodium dodecyl sulfate (SDS) solution and rapidly mixing. The sticky mixture was kept on ice for 2 min, after which 3 ml of phenol mix was added (phenol, saturated with 0.01 M Tris-HCl buffer, pH 7.6, 10% (w/v) with respect to m-cresol and 0.5% with respect to 8-hydroxyquinoline). The emulsion was shaken for 10 min on a wrist-action shaker in a 4°C cold room, and the phases were separated by centrifugation at 12,OOOg for 10 min. The aqueous phase was removed and transferred to 3 ml of 241 chloroform-isoamyl alcohol kept in ice. ARer shaking rapidly for 2 min, the aqueous phase was removed and mixed with 6 ml of 95% ethanol kept at -20°C. Precipitation of RNA was allowed to proceed overnight. RNA was recovered by centrifugation of the precipitate at 12,OOOg for 10 min. The ethanol was discarded, and the RNA was dissolved in 0.1-0.3 ml of buffer II (0.01 M Tris-HCl, pH 7.6, 0.1 M sodium acetate). The concentration of the RNA solution was about 20 OD units/ml. In order to facilitate the application to the gel, it was necessary to add a drop of 40% sucrose in buffer II, containing enough bromophenol blue to color the final solution. Isolation of cytoplasmic ribosomes. Ribosomes were isolated by a modification of the method of Rawson and Stutz (8). All manipulations were carried out at 4°C. Two hundred milliliters of resting cells were washed in buffer III (0.01 M Tris-HCl, pH 7.6, 0.004 M MgCl,, and 0.04 M KC11 by centrifugation at SOOg for 5 min. The cells were resuspended in 7 ml of buffer III and disrupted in a French press (3000 psi), followed by centrifugation at 15,000 rpm for 10 min in a Sorvall SS-34 rotor. The ribosomes were isolated by pelleting through 2 ml containing 1.5 M sucrose in 0.01 M Tris-HCl, pH 7.6, 0.001 M MgCl,, and 0.01 M KC1 at 36,000 rpm for 3.5 h in a Spinco 40 rotor. The ribosome pellet was resuspended in buffer IV (0.001 M Tris, pH 7.6, 0.001 M MgCl,, and 0.01 M KC11 at a concentration of about 1

rRNA

TRANSCRIPTION

203

mg/ml, and the suspension was clarified by centrifugation at 10,000 rpm for 5 min in a Sorvall SS-34 rotor. About 0.5 ml (0.5 mg) was layered on a 1530% (w/v) linear sucrose gradient made in buffer IV and centrifuged in a Spinco L-2 ultracentrifuge in an SW 27 rotor at 27,000 rpm for 5 h at 3°C. The gradient was analyzed using the flow-through cell assembly of No11 (9) in a Gilford Model 203 spectrophotometer, and the fractions were collected. The fraction containing the ribosomal peak was centrifuged at 36,000 rpm for 3 h in the Spinco 40 rotor. The supernatant was discarded and the pellet was frozen at -20°C. Isolation of polysomes. Two hundred milliliters of cells were washed and resuspended in 5 ml of buffer III made 20% (w/v) with respect to sucrose and were disrupted in a French pressure cell at 1000 psi followed by a centrifugation at 10,000 rpm for 10 min in an SS-34 Sorvall rotor at 3°C. An aliquot (0.5 ml) was layered on a 7-40% (w/v) surcose gradient made in buffer III and was centrifuged at 27,000 rpm for 3 h in a Spinco SW 27 rotor at 3°C. Fractions were collected from the gradients, and the polysome fractions were combined and ribosomes were recovered by centrifugation at 25,000 rpm for 5 h. Isolation of cytoplasmic ribosomal subunits. The ribosome pellet was resuspended in buffer IV made 0.02 M with respect to EDTA. About 0.5 ml of a 1 mg/ ml suspension was layered on a lo-30% (w/v) sucrose gradient in buffer IV made 0.02 M with respect to EDTA and was centrifuged at 27,000 rpm for 10 h in a Spinco SW 27 rotor at 3°C. Fractions were collected as described before and were frozen after pelleting. Extraction of RNA from ribosomes, ribosomal subunits, and polysomes. The ribosomal pellet was resuspended in 1 ml of buffer II and 0.1 ml of SDS (20%, w/v) was added. The mixture was warmed to room temperature for about 1 min and immediately chilled. Two milliliters of buffer II were added, followed by 6 ml of cold 95% ethanol. RNA precipitation was allowed to occur overnight. Gel preparation. The method used was essentially that of Peacock and Dingman (10) employing 2.7% acrylamide (cyanogum 41, EC Corp.) gels containing 0.5% agarose (Seakern) polymerized with ammonium persulfate (EC Corp.) in an EC slab-electrophoresis apparatus. After a l-h prerun at 200 V, the RNA sample of 0.025 ml, containing 1.5-2.0 A,,,, units was applied, and electrophoresis was carried out at a current of 40-45 mA until the tracking dye had travelled about three-fourths of the length of the gel slab. The gels were stained with a solution containing 0.2 M sodium acetate, 0.2 M acetic acid, 0.4% methylene blue, pH 4.7, for lo-15 min and were then destained overnight in running water. The bands were recorded using a Joyce-Loebl recording microdensitometer. The proportionality of the final

204

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recording to the RNA concentration was verified in control experiments. For measurement of radioactivity the bands were cut from the gels with a stack of parallel razor blades, and each slice was placed in a scintillation vial with 6.0 ml of scintillation fluid (100 g of naphthalene and 5 g of 2,5diphenyloxazole in a liter of 1,4-dioxane), and the vial was counted in a Beckman LX50 scintillation counter. The specific activities of RNA recovered after electrophoresis, from different amounts of labeled RNA applied to the gels, were found to be the same, verifying the proportionality of the gel counting method. Cell counts. Cell counts were done using a Coulter counter (Model A1 (11). Samples of the cultures were usually diluted 1:50 with Millipore-filtered 0.4% NaCl containing 0.01% merthiolate. Chlorophyll determinations. Five to ten milliliters of cell cultures were centrifuged. A pinch of MgCO, was added to the pellet followed by 2 to 5 ml of acetone. The sample was left overnight in the dark and was then centrifuged. The absorption of the supernatant was examined in a Cary (Model 141 spectrophotometer at 663 and 645 nm, and the amounts of chlorophylls a and b were estimated as described previously (12). RESULTS

A major separation

AND

DISCUSSION

problem in the extraction and of the cytoplasmic rRNAs of

SCHIFF

has been their instability which leads to the formation of a 22s component from the 25s rRNA. Since the plastid rRNA has a component at 23S, it was very important to be able to distinguish these two species and to control the degradation of the 25s RNA. Heizman (7) found that high concentrations of sodium thioglycollate prevented the degradation of 25s rRNA to 22S, and Rawson et al. (13) showed that sodium ions, regardless of the anion, achieved the same protection of the 25s rRNA.

Euglena

Integrity of RNAs Purification

during

Extraction

and

Figure 1 shows that RNA extracted from mutant W,BUL in which plastid DNA and plastid rRNA are undetectable still exhibits a small band at 22s indicating a small amount of degradation even when sodium thioglycollate is present. When rRNAs are extracted from fully green wild-type cells (Fig. 11, using the same techniques, the 22s degradation product of the 25s cytoplasmic rRNA can be sepa-

DIRECTION OF MOVEMENT -

c

FIG. 1. Total RNA extracted from W,BUL cells. RNA was extracted and separated on gel electrophoresis as described in Materials and Methods, stained, and scanned. The inset shows rRNA from wild-type cells extracted with different concentrations of sodium thioglycollate showing the appearance of the 22s degradation product and its separation from 23s plastid rRNA. Light-grown cells rested in the light were resuspended in buffer I supplemented with 0.25, 0.1, and 0.05 N sodium thioglycollate, and RNA was extracted as described in Materials and Methods.

PHOTOREGULATION

OF

rRNA

TRANSCRIPTION

205

rated from the 23s chloroplast rRNA com- collate was omitted (data not shown) they ponent. When sodium thioglycollate is re- are not degradation products of the cytoplasmic rRNAs. A 7S fraction was reduced in concentration to allow degradation of the 2% RNA to occur, the 22s com- ported in other systems (16) which could be ponent increases at the expense of the 25S, released from the large ribosomal subunit by heat, urea, or dimethylsulfoxide treatbut the constant 23s component remains. Thus, the degradation of the 2% can be ment and was thought to originate from kept low using sodium thioglycollate dur- the cleavage of the precursor during rRNA Upon separation of the subing extraction, and this small amount of maturation. units of the cytoplasmic ribosomes of Eudegradation product can easily be distinguished from the 23s plastid ribosomal glena using EDTA (data not shown), most rRNA. In our hands, the chloroplast riboof the 5 and 7s RNA is found to be associated with the large 60s subunit as it is in somal rRNAs behave as 23 and 16s entities on gels, show coelectrophoresis with other eukaryotic systems (16). the corresponding rRNAs from Eschein Dark-Grown Wild Type and richia coli and have molecular weights of pl-rRNA WguL 1.2 and 0.56 x 106, respectively, in good agreement with the values found by other Figure 2 shows that we can routinely workers (6, 14). The cytoplasmic rRNAs detect bands in RNA preparations from from wild type and W,BUL show the same dark-grown wild-type cells which correvalues of 25 and 20s which correspond to spond to the 23 and 16s pl-rRNA compomolecular weights of 1.4 and 0.88, respec- nents. The 23s band is clearly separated tively, and are also in agreement with from the small 22s component representthese reports. ing the degradation product of the cytBased on studies of the degradation of rRNA. Repeated electrophoresis of the the 25s cytoplasmic rRNA in the absence RNA from W,BUL, a mutant lacking deof protective sodium ions and the fact that tectable plastid DNA, fails to reveal any 23 there is a stoichiometric relation between or 16s components and only shows the 22s the 25s that disappears and the 22s which and other degradation products of the cytappears, Heizman (7) proposed that a nu- rRNAs. These results are in agreement clease was tightly bound to the 25s RNA with the expectation that W3BUL should which nicked it in a specific spot leading to the observed degradation and that sodium thioglycollate inhibited the action of this enzyme. We have found, however, that the same pattern of degradation can be obtained if the RNA is subjected to shearing forces in a French pressure cell in the presence of sodium thioglycollate. This result suggests that the 25s rRNA has sites which can be disrupted mechanically in a reproducible fashion. Of course, it is possible that these sites are created through prior nuclease action. Figure 1 also shows that 35 and 32s components can be detected which have molecular weights of 3.1 and 2.4 x 106, respectively, and correspond to the cyto8 DIRECTION OF MOVEMENT c 3 plasmic ribosomal RNA precursors (14). Also displayed are the 4S tRNAs and the 5 FIG. 2. Comparison of rRNA from dark-grown, and 7S reported for other eukaryotic riboW,BUL and wild-type cells. Total cellular RNA was somes (15-17). Since the 5 and 7S compoextracted, separated on acrylamide, stained, and scanned as described in Materials and Methods. nents did not change when sodium thiogly-

206

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AND

WHIFF

lack pl-rRNAs since it lacks plastid DNA, and it has been shown that the pl-rRNA cistrons are located exclusively in the plastid DNA (46, 47). Wild-type dark-grown cells contain plastid DNA, and plastid ribosomes are clearly evident in the proplastids of dark-grown cells by electron microscopy (181, in agreement with the presence of pl-rRNAs in these cells (Fig. 2). Two laboratories (14, 19) have reported the absence of pl-rRNAs from dark-grown Euglena cells, but it is not clear whether the techniques employed or the strain differences between bacilluris and Z (which they employed) prevented their detection. Constancy of Total DNA plast Development

during

Chloro-

Since the total amounts of RNA recovered from cells was different from one experiment to another it proved advisable to have an internal standard to which the OD and radioactivity of the isolated rRNAs could be compared. Since the total DNA of the cells was extracted along with the RNAs and formed a discrete peak on the acrylamide gels, the amount of DNA on the gels was used as an internal standard with which the RNAs were compared. We were able to verify that the ratio of RNA recovered to DNA recovered was constant over a wide range of cell densities, using a single batch of cells at various cell concentrations. We also verified that there was no appreciable synthesis of DNA during chloroplast development since the amount of 32Pi incorporated into DNA was negligible under conditions where the RNAs were highly labeled, which would be expected since nondividing cells are used in all experiments. Changes in rRNA Peaks during LightInduced Chloroplast Development in Resting Cells Figures 3 and 4 show that the concentrations of 25 and 20s cyt-rRNAs and the small amount of degradation product of the 25S, 22S, remain fairly constant, relative to DNA, during chloroplast development. The 23 and 16s pl-rRNAs increase X-fold during plastid development, and by

DIRECTION

OF MOVEMENT

-

1

FIG. 3. The accumulation of 23 and 16s pl-rRNA during light-induced chloroplast development in dark-grown wild-type resting cells. Dark-grown resting cells were exposed t,o light and 25-ml samples were withdrawn every 12 h. RNA was extracted and separated by gel electrophoresis, stained, and scanned as described in Materials and Methods. Since recoveries and amounts applied to the gels vary, compare 23 and 16s pl-rRNAs with 25 and 20s cyt-rRNAs or with internal DNA which do not change during development.

48 h of development the maximum increase is obtained, representing 25% of the cellular rRNA. Similar values were obtained in bacilluris (11, 45) and in the Z strain (7, 44), but a maximum of 7% was reported during chloroplast development in resting cells of the Z strain (19). Linear extrapolation of the values for pl-rRNAs back to zero time yields a value of 2% of the total rRNA of the cell for the pl-rRNA of the proplastids of bacillaris in the present work.

9

c o

0

0

PHOTOREGULATION

OF

25S+2OS DNA 20

000

25 S (cprdpg) (LIGHT)

0

12 24 36 48 60 DURATION OF LIGHT

72 84 (HRS.)

FIG. 4. A comparison between the changes in relative amounts of pl-rRNA and cyt-rRNA during light-induced chloroplast development in resting cells and the labeling of these species by 12-h pulses of ,“Pi during this period. The relative changes in the amounts were obtained from the data of Fig. 3. $>PI was introduced into cultures of dark-grown resting cells either kept in the dark or exposed to light for the appropriate time, at 12-h intervals, and each culture was extracted after 12 h of labeling. RNA was extracted, separated, and counted as described in Materials and Methods.

Labeling of rRNAs during Light-Induced Chloroplast Development in Resting Cells As may be seen in Fig. 4, 32Pi, administered as 12-h pulses to dark-grown resting cells after the time of exposure to light, is incorporated into both cyt-rRNAs and plrRNAs as previously observed (11). The labeling of the pl-rRNAs represents net synthesis since the amounts of these rRNAs increase during plastid development. The labeling of the cyt-rRNAs, however, represents turnover since there is no increase in the amount of cyt-rRNA during development. The amount of 32Pi incorporated into both cyt-rRNA and pl-rRNA is quite similar but Fig. 4 shows that the specific activ-

rRNA

TRANSCRIPTION

207

ity of pl-rRNA is three to four times higher than cyt-rRNA, as also found in previous experiments (11). This difference can be traced to the fact that the plastid enters development with a minute amount of nonradioactive rRNA whose contribution to the specific activity is very small and the specific activity at the end of development is, therefore, high. The cytoplasm, however, contains a large amount of preexisting nonradioactive rRNA which results in a lower specific activity for a comparable amount of synthesis. Thus a comparison of the specific activities of the cyt-rRNA and pl-rRNA does not give a true indication of the relative rates of synthesis. In comparing the kinetics of synthesis of rRNA in the chloroplast and the cytoplasm, therefore, we cannot use the specific activity value of pl-rRNA since it is constant after 24 h. Therefore, the relative amount of pl-rRNA, represented by the value calculated from the OD is used directly. On the other hand, the relative amount of cyt-rRNA remains constant during development, and therefore the specific activity of this fraction is the meaningful value to be considered. Figure 4 shows that pl-rRNA begins to increase immediately upon illumination and continues at a constant rate for about 48 h, when synthesis ceases. Brown and Haselkorn (141 presented data showing that by 24 h pl-rRNA synthesis is completed in the Z strain, although a careful examination of their data indicates that synthesis might in fact continue up to 74 h. Scott et al. (19) reported that the full amount is synthesized during the first 16 h in the Z strain, but in this case pl-rRNA comprised only 7% of total cellular RNA. It is interesting that pl-rRNA begins to accumulate immediately without the lag of 12 h which is usually observed for other chloroplast constituents, e.g., cytochrome 552 and ribulose-1,5-diphosphate carboxylase (20). These enzymes are synthesized, at least in part, on chloroplast ribosomes, and the low amounts of chloroplast ribosomes in the proplastids might be a limiting factor in the appearance of these enzymes.

208

COHEN

AND

In contrast chloroplast enzymes which are nuclear coded and synthesized on cytoplasmic ribosomes increase without a lag (20).

Figure 5 shows that synthesis of cytrRNA begins at the instant of illumination with the rate slowly increasing in the first 12 h; a higher rate is achieved between 12 and 36 h. This accompanies the increase of many cytoplasmically synthesized plastid proteins (20-22). The pulse experiment in Fig. 4 reveals that the rate of turnover decreases abruptly after 36 h and then continues at the same rate as found in darkgrown cells. The fact that dark-grown resting cells and resting cells containing fully formed chloroplasts turn over their cytrRNA at the same low rate indicates that this suffices for the formation of the minimum rRNA necessary for the limited protein synthesis in these starving cells. Figure 5 shows that the specific activity of pl-rRNA reaches a constant value 24 h after illumination while, according to the absorbance values (Fig. 4), synthesis continues linearly for 48 h. This can be understood in the following way: pl-rRNA is synthesized rapidly upon illumination. The specific activity of the newly made rRNA reflects the specific activity of the available phosphate and nucleotide (Pi-N) pools z 24

23s pl-rRNA

B g

x

26

X

X X

:

25s cyt -rRNA WILD 0

0

24 TIME

TYPE 0

IN LIGHZ 0

72 IN LIG?

(HOURS)

FIG. 5. 32Pi incorporation into 25s cyt-rRNA and 235 pl-rRNA of dark-grown resting wild-type and 255 cyt-rRNA of W,BUL during 72 h of illumination. Dark-grown resting cells were exposed to light in the presence of 32Pi. A 25-ml sample was withdrawn every 12 h for RNA extraction, separation, and determination of RNA and radioactivity.

SCHIFF

in the cells. Since the amount of unlabeled pl-rRNA present in dark-grown cells is negligible, its contribution is diluted out by the increasing amount of newly labeled pl-rRNA. The specific activity of the total pl-rRNA, after sufficient dilution has taken place, would almost equal the specific activity of the phosphate pool and would therefore be constant. Estimation of changes in amounts can be obtained only by considering the relative absorbances of the pl-rRNA peaks (Fig. 4). In the cytoplasm the situation is reversed. Here, the amount of cyt-rRNA measured as relative absorbance of the cytoplasmic peak is constant. This peak contains both the preexisting, nonradioactive RNA and the newly synthesized material. A comparison of the specific activities at different points during development is meaningful in this case and represents the relative amount of the newly made cyt-rRNA. The actual amount, however, is difficult to calculate since there is no net accumulation of cytrRNA corresponding to the radioactivity incorporated. The following analysis is made to obtain a rough estimate of the extent of synthesis or turnover of cytrRNA. The fact that the net accumulated plrRNA is of constant specific activity means that the specific activity (sp act) of plrRNA equals sp act of the Pi-N pools in the chloroplast. The isotope in the cytoplasm and in the chloroplast probably equilibrates rapidly, although the size of the pools need not be the same. Therefore, the sp act of the chloroplast Pi-N pool equals sp act of the cytoplasmic Pi-N pool, and the sp act of pl-rRNA = sp act of the cytoplasmic Pi-N pool. The sp act of the newly made cyt-rRNA obviously reflects the sp act of the cytoplasmic Pi-N pool from which it is made. The amount of this RNA can therefore be calculated from the fact that the sp act of the newly synthesized cyt-rRNA equals sp act of the pl-rRNA. To make this calculation, the sp act of the pl-rRNA and the counts incorporated into the cyt-rRNA peak were taken from the same experimental sample at 36 h of development when the specific activity of pl-rRNA has become constant.

PHOTOREGULATION

OF

Table I shows the results of these calculations; the radioactivity incorporated into the cyt-rRNA peak represents 60% of its total OD. Since net synthesis is not observed, we are forced to the conclusion that there is extensive turnover of the cytrRNA and, as a result, approximately 60% of the RNA has been degraded and resynthesized during 36 h of illumination. A similar phenomenon has been observed in resting liver tissue (23), in contact-inhibited lymphocytes (24), and in slime mold during morphogenesis, where 75% of the ribosomes are turned over (25). The total RNA content of the resting Englena cells does not change significantly during chloroplast development (11, 361, and the amount of cyt-rRNA does not change (Fig. 4). Since the amount of plrRNA which is synthesized during chloroplast development represents 25% of the total rRNA, the nucleotides to form this RNA cannot come from the degradation of cyt-rRNA and must be provided by the degradation of some other species of RNA or from the appreciable preexisting pools of nucleotides known to exist in Euglena cells (26). Labeling

of Cyt-rRNA

in W,@UL

It was of interest to determine whether the light induction of cyt-rRNA observed upon illumination of dark-grown resting wild-type cells could be reproduced in mutant WBBUL which lacks detectable plastid DNA and has little or no plastid strucTABLE THE

AMOUNT

DURING

Zperiment

1 2

36

OF 25s HOURS

cyt-rRNA

OF WILD-TYPE

OF LIGHT-INDUCED

CELLS

CHLOROPLAST

rRNA

209

TRANSCRIPTION

ture. Figure 5 shows that the cyt-rRNA of W,BUL becomes labeled when resting cells are illuminated in the presence of 32Pi, but the rate and final extent of labeling are lower than in wild type. It appears that W3BUL retains a cytoplasmic component of light induction similar to wild-type but the elimination of the plastid photoreceptor system and most plastid structures do not allow the full induction to take place. It is possible that the synthesis of cyt-rRNA can only be fully induced by light when a normal proplastid is present to serve as a sink for the products of cytoplasmic protein synthesis or that a signal must be sent from the developing plastid to sustain a high level of synthesis in the cytoplasm (36). The induction of cyt-rRNA synthesis in W3BUL is consistent with the finding that the cistrons for this RNA are coded in nuclear DNA (46,47). Chloroplast constituents which are not coded in chloroplast DNA and are probably coded in nuclear DNA include the NADP-triosephosphate dehydrogenase (201, alkaline DNase (43), leucyl-tRNA synthetase (28), and the photoreactivating enzyme (29). Of these, the first two cannot be induced by light to increase in resting cells of W3BUL. The photoreactivating enzyme, however, like the labeling of the cyt-rRNA can be induced to increase in W,BUL (29). These activities join paramylum degradation (30) and the formation of reducing power for chloramphenicol detoxification (31) as examples of activities that are light inducible in W,BUL. I SYNTHESIZED DEVELOPMENT

AND

DEGRADED

THROUGH

IN DARK-GROWN

RESTING

TURNOVER CELLS”

Counts incorporated into 25s cyt-rRNA in 36 h (cpm)

Total amount of 25s cytrRNA (OD)

Specific activity of 23s RNA at 36 h (cpm/OD unit)

Amount of 2% RNA represented bv the counts incorporated (OD)

Percenta e of total 25s fi NA represented by newlv svnthesized”253 RNA by 36 h

1.35 x 105 1.82 x lo”

9.2 10.4

2.4 x lo4 2.9 x 104

5.6 6.2

60 58

n The experimental values were obtained from the experiment described in Figs. 7 and 8. The calculation was based on the assumption that the specific activity of the phosphate pools in the chloroplast and in the cytoplasm are the same when the pl-rRNA has reached constant specific activity and, therefore, the specific activity of the newly made pl-rRNA equals the specific activity of the newly made cyt-rRNA which together with the preexisting unlabeled cyt-rRNA compose the total cyt-rRNA peak observed.

210

COHEN

AND

Labeling of the rRNA of Cytoplasmic Polysomes on Light Induction of Plastid Development

Figure 6 shows that, when 32Pi is administered to dark-grown resting cells during the first 3 h of light-induced chloroplast development, the rRNA of the cytoplasmic polysomes has a fourfold higher specific activity than the rRNA of the cytoplasmic monosomes. This indicates that the newly synthesized cyt-rRNA is used to form ribosomes which immediately become associated with polysomes and that the label enters the monosome pool through dissociation of the polysomes into monosomes after a round of protein synthesis is completed. It is possible that existing monosomes in the- starved dark-grown-resting cells cannot be used for protein synthesis and that messenger RNA formed on light induction only binds to newly synthesized monosomes explaining why turnover of cyt-rRNA takes place upon light induction of chloroplast development. Consistent with this explanation, Heizmann et al. (32) showed that most of the cytoplasmic ribosomes in resting cells are found in the monosome fraction. The monosome frac-

SCHIFF

tion of HeLa cells has been reported (33) to be inactive in protein synthesis, and Kaempfer (34) has proposed that the number of ribosomes active in protein synthesis in E. coli is controlled by allowing excess subunits to form inactive monosomes. The Light Dependence of rRNA in Wild-Type Resting Cells

Labeling

Figure 7 shows the results of experiments which demonstrate that the control of formation of cyt-rRNA and pl-rRNA are different. Both rRNAs become labeled upon light exposure, and this labeling continues in uninterrupted illumination. However, when the dark-grown resting cells are given a brief pulse of light fol0

6

3

6

9

12

15

x0 X

4- 25s cyt-rRNA

4

J

E 3-

z 5 12- 23s pl-rRNA I= s _ E9

cl x 0 0

ti

/

6-

%I

/”

3-

/: B 0

0

4

8 VOLUME

12 16 COLLECTED

20 (ml1

24

28

FIG. 6. The light-induced labeling by 32P, of the polysomal and monosomal fractions of W,BUL and the specific activities of their rRNAs. Fifty milliliters of dark-grown resting wild-type cells were exposed to light for 3 h in the presence of 32Pi, and polysomes were extracted and separated on a sucrose gradient as shown. Fractions were collected and rRNA was extracted, separated, and counted as described in Materials and Methods.

3

I I 6 9 12 TIME (HOURS)

I 15

FIG. 7. The synthesis of 25s cyt-rRNA and 23s pl-rRNA in dark-grown resting wild-type cells in continuous light and in cells illuminated for 90 min, returned to darkness for 10 h, and then reilluminated. A dark-grown wild-type resting culture was divided in half. 32Pi was added to both, and the cultures were illuminated. One culture was returned to darkness after 90 mm and reilluminated after 10 h of darkness. RNA was extracted and separated at the times indicated, and the amount and radioactivity was determined.

lowed by darkness, it is evident that the labeling of cyt-rRNA is strictly light dependent while that of the pl-rRNA is not since labeling of cyt-rRNA ceases in darkness while pl-rRNA labeling continues in darkness after a light pulse. It is known from other experiments (35) that when dark-grown resting cells of Euglena are given a l-2-h pulse of light and are returned to darkness for 12 h, upon reillumination the usual 12-h lag in chlorophyll synthesis is eliminated; it appears that events which ordinarily take place in the lag period of continuous illumination can take place in complete darkness after a brief pulse of light. One of these events might be the transcription of pl-rRNA. It is also known that the increase in several plastid enzymes which are synthesized outside the chloroplast is strictly light dependent during normal plastid development in continuous light and that these fail to increase in the dark period after a pulse of light, consistent with the finding reported here that transcription of cytrRNA is strictly light dependent (20, 39). The Synthesis and Decay of cyt-rRNA beling in Resting Cells of W$lJL

La-

In order to study the strict light requirement for cyt-rRNA labeling we turned to mutant W,BUL in which chloroplast DNA, pl-rRNA, most proplastid structures, and the protochlorophyll(ide) photocontrol system of the plastid are undetectable (18, 42). We have already shown that the labeling of cyt-rRNA can be induced in W:%BUL, although to a lesser extent than in wild type (Fig. 5). Dark-grown resting cells of W3BUL were exposed to light for various times less than 3 h in the presence of “*Pi and were then returned to darkness for the remainder of the 3-h period, at which point RNA was extracted. Figure 8 shows that cells exposed to 45 min of light and then returned to darkness incorporated as much “Pi into rRNA as cells exposed to light for the full 3-h period. The time in light is limiting below 45 min, but exposure for longer times does not yield additional labeling. Continuous light, then, is not necessary to achieve maximum synthesis, in-

dicating a possible trigger mechanism. Previous experiments (Fig. 7) have shown that the synthesis in cells returned to darkness is not sustained beyond the first 3 h, and the ability to incorporate “‘Pi into rRNA at the same rate as cells in continuous light soon decays. This was examined in the next experiment. Cells were exposed to light for 45 min and returned to darkness. The rate of synthesis in the dark was examined in 3-h pulses ending at the ninth hour. A 45-min light period was chosen since it produced the same response as in cells in continuous light for the first 3 h. Figure 9 shows that during the remainder of the first 3 h there was no decay (see also Fig. 81, but the rate of labeling decreased with a half-life of about 3 h thereafter in the dark. It is possible, of course, that 45 min of illumination is optimal only for the synthesis which occurs over the remainder of the first 3-h dark period and not thereafter but that light given for longer periods might delay the decay for longer periods in darkness. Figure 10 confirms that prolongation of the light period beyond 45 min allows increased synthesis to occur for the remainder of a 6-h dark period.

Y-

0

30

60 TIME IN LIGHT

90 (MIN.)

IS0

1

FIG. 8. The time in light required to achieve as much synthesis of cyt-rRNA in W,BUL cells as in comparable cells in continuous light. Dark-grown resting cells were illuminated for the times indicated and then returned to darkness for the remainder of a 3-h period. 32P, was present for the entire 3 h. RNA was extracted at the end of the 3-h period, separated, and estimated and counted as described in Materials and Methods.

212

COHEN 075

3

6

9

AND

12

9

TIME

SCHIFF

scription in the cytoplasm and plastid, broad band action spectra were measured. Figures 11 and 12 show that the labeling of cyt-rRNA in W,BUL and wild type saturates at about 100 pW/cm2 which is consistent with other light saturation curves for plastid development in Euglena (5). Labeling of pl-rRNA also saturates at this intensity in wild-type cells (Fig. 12). Saturation curves in broad band monochromatic light for labeling of rRNA (Fig. 13) show that blue light is about as effec-

(HOURS)

FIG. 9. Comparison of the rate of synthesis of 255 cyt-rRNA in W,BUL resting cells in continuous light and in cells illuminated for 90 min and returned to darkness. Cells were rested in eight flasks. At zero time, seven flasks were illuminated and three were returned to darkness after 90 min. 32Pi was administered at 3-h intervals to different cultures, and each culture was extracted after being exposed to 3zPi for 3 h. The incorporation, therefore, represents the amount incorporated over the 3-h interval indicated for each set of conditions.

1

I 0

It appears that the control system which makes the transcription of cyt-rRNA strictly dependent on light has a finite decay time in darkness which is dependent on the length of the illumination period. Forty-five minutes of illumination is sufficient to allow maximal synthesis for the next 3 h in darkness but, after this time, a longer illumination period is necessary to prevent decay. Whatever the mechanism, a factor or factors with a decay time of about 3 h seem to be involved. The Spectral Dependence Cyt-rRNA and pl-rRNA and W,BUL

TI&

IN LIGHT

4 (HOURS)

6

FIG. 10. “*Pi incorporation into 25s and cytrRNA of W,BUL during a pulse of 52P, lasting 3-6 h after the cells were illuminated for various times and returned to darkness. Dark-grown resting W,BUL cells were exposed to light for the times indicated and were then returned to darkness. They received “*Pi 3 h after the beginning of illumination for a 3-h period in all cases and were then immediately extracted.

of Labeling of in Wild Type

At least two photoreceptor systems are thought to be involved in the induction of plastid development in Euglena by light. One of these is the protochlorophyll(ide) receptor system of the plastid which absorbs in blue and red light and the other is a nonplastidic receptor system which absorbs mainly in the blue region of the spectrum (18, 27, 30, 31, 35). In order to learn which photoreceptors control rRNA tran-

A’ INTENSITY

(pWATTS/cm*l

FIG. 11. The labeling of the 2% cyt-rRNA of W,BUL as a function of light intensity. Dark-grown resting W,BUL cells were exposed to various intensities of white light for 3 h in the presence of 32Pi (0.05 mCi/ml). The high intensities, above 600 fc, were obtained using incandescent spotlights, and the values were normalized to the values obtained using fluorescent light.

PHOTOREGULATION

OF

II

23: rRNA

50r--x

I

:’

40. /

I00

200 300 INTENSITY (~WATTS/cm21

600

FIG. 12. The labeling of the 25s cyt-rRNA and 235 pl-rRNA of wild-type cells as a function of light intensity. Dark-grown resting cells were exposed to various intensities of white light in the presence of 32Pi for 3 h after which rRNA was extracted.

1W,BUL 25s cyl-rRNA El-

cl ; 2

WILD TYPE 25s cyf-rRNA

rRNA

213

TRANSCRIPTION

tive as white light as an inducer of cytrRNA labeling in wild-type and W,BUL cells while green and red light are quite ineffective. The labeling of pl-rRNA in wild-type cells, however, is induced best by blue light which is as effective as white, but red light is also quite effective although less than blue, while green is still less effective. Table II shows that the ratio of effectiveness of blue to green to red corresponds well with the absorption ratios of the protochlorophyllide holochrome from beans. These results indicate that the transcription of cyt-rRNA is under control of the nonplastid blue light receptor system while the transcription of pl-rRNA is controlled by the protochlorophyll(ide) system of the plastid. Among the processes known to be controlled by the blue light receptor in Euglena are lag elimination by preillumination (27), paramylum degradation (301, and mobilization of reducing power for chloramphenicol reduction (31). A high molecular weight photoreversible pigment has recently been described in Euglena extracts which is an excellent candidate for the blue light receptor (37) and is very similar to a similar pigment isolated from Dictyostelium (38). Action spectra which TABLE

LL Y c%

II

RELATIVE EFFECTIVENESS OF THREE SPECTRAL REGIONS IN PROMOTING pl-rRNA SYNTHESIS COMPARED WITH THE RELATIVE ABSORPTION OF THE PROTOCHLOROPHYLL(IDE) HOLOCHROME WILD TYPE

100 150 200 50 INTENSITY bWATTS/cm’)

FIG. 13. Effectiveness of the various spectral regions in promoting the labeling of rRNAs of W,BUL and wild type. Dark-grown resting cells were exposed to increasing intensities of the three spectral regions for 3 h in the presence of ?‘Pi; RNA was then extracted. (X-X), White light; (O-O), blue, 400510 nm; (B-m), red, 610-700 nm; (A-A), green, 520-570 nm.

Spectral

region

Blue (400-510 nm) Green (520-570

Energy required for 50% maximum incorporation (wW/ cmy)

Relative --------Observed

effectiveness (1 Expetted

18

7.4

6.0

140

1.0

1.0

55

2.5

3.0

b

nm) Red (610-700 nm)

o Red and blue were normalized to green taken as 1.0 for both observed and expected values. * The expected effectiveness of the protochlorophyll(ide) holochrome was obtained from Holowinsky and Schiff (35).

214

COHEN

AND

indicate involvement of a blue light receptor system in plastid development have also been reported in Cyanidium (40) and in a mutant of Scenedesmus (41). Among the processes known to be controlled by the protochlorophyll(ide) system in Euglena are the synthesis of chlorophyll and of two plastid enzymes synthesized outside the chloroplast, NADP-triosephosphate dehydrogenase and alkaline DNase (27). CONCLUSIONS

We ordinarily study the light-induced development of chloroplasts from proplastids in dark-grown resting cells of Euglena where cell division has been inhibited by nutritional deprivation to allow the study of plastid development without variables due to cell division. Turnover of proteins and RNA is consequently quite high in these cells as in other starving microorganisms in order to provide the constituents necessary for plastid development. A particular example of this turnover is found in the behavior of the cyt-rRNA upon light induction of plastid development in resting cells where 60% of this RNA is turned over during the course of chloroplast development but the total amount per cell does not change. This turnover, however, probably does not exist to provide precursors for the formation of pl-rRNA which takes place by net synthesis of an amount which constitutes 25% of the total rRNA of the cells by the end of chloroplast development. Since the total cellular RNA remains fairly constant over the course of development (361, a decrease of 25% in the cyt-rRNA or an amount approaching this in the total cellular RNA would be expected if cyt-rRNA were sacrificed to form pl-rRNA. A more likely explanation for these findings is that in the dark-grown resting cells most of the cytoplasmic ribosomes exist as monosomes. Upon light induction of chloroplast development, polysomes are required to synthesize the proteins contributed by the cytoplasm to the chloroplast. Since we find that the rRNA of the cytoplasmic polysomes becomes much more highly labeled than the monosomes, we infer that the newly synthesized rRNA enters the ribosomes of the polysomes first and these

SCHIFF

eventually find their way into the monosome fraction after completing protein synthesis. Thus the cell might be incapable of activating preexisting monosomes as in other systems but must synthesize new ribosomes in order to enable them to participate in new protein synthesis associated with chloroplast development. This would explain the need for turnover of cytrRNA since only newly synthesized ribosomes can apparently be programmed for protein synthesis. The net synthesis of an appreciable amount of pl-rRNA is not unexpected since the proplastid undergoes an increase in volume of about 60-fold in becoming a chloroplast and additional ribosomes must be provided for the greater demands of the larger structure (18). The synthesis of both cytoplasmic and chloroplastic rRNAs reach their maximum rates soon after light induction and remain high until about 36 h of development. Thus the protein-synthesizing machinery is formed well ahead of the time of maximal protein synthesis which is about 15 h in the case of proteins supplied by the cytoplasm to the developing plastid and about 36 h for processes taking place within the developing plastid (18, 20). In order to have the maximal induction of cyt-rRNA synthesis, a proplastid must be present since light induction of cyt-rRNA in W,BUL, a mutant in which plastid DNA, the protochlorophyll(ide) light control system, and many plastid structures are undetectable, occurs to a much lower extent than in wild type (see also 11,361. The fact that it occurs at all suggests that a photoreceptor system other than the protochlorophyll(ide) system must exist in the nonplastid compartments of the cell. At least two photoreceptor systems are present in Euglena (l&27,37): a blue light system which controls the nonplastid contributions to chloroplast development and the red- and blue-absorbing protochlorophyll(ide) system within the plastid which controls plastid-localized events including chlorophyll synthesis. The labeling of cytrRNA is strictly light dependent during the first 12 h of greening, and its action spectrum in both wild type and W,BUL indicates that it is under the control of the

PHOTOREGULATION

OF

blue light receptor system. The formation of pl-rRNA in wild type, on the other hand, is induced by a brief illumination and then continues in darkness; the action spectrum indicates that it is under control of the protochlorophyll(ide) system of the plastid. Thus the formation of chloroplastic and nonchloroplastic rRNAs are regulated quite differently from each other during plastid development. This is not unexpected since the formation of a chloroplast requires contributions from inside and outside the plastid and these must be properly coordinated if development is to proceed normally. Although the broad band action spectra reported here are consistent with the detailed blue light system and protochlorophyll(ide) action spectra reported previously (27) and with the absorptions of pigments which can be isolated from the cells (4, 371, there is little information as to where these pigments act in regulating plastid development. Of course it is possible that they act at the substrate level, as protochlorophyll(ide) does in serving as a precursor which is photoreduced to chlorophyll, and that any regulatory effects on ot’her processes such as transcription of rRNAs is a secondary effect of the substrates formed or removed in these photoreactions. Another explanation is possible, however. The entire process of light-induced chloroplast development in Euglena resembles substrate induction of enzymes on a large scale. While substrate induction in many microorganisms involves the formation of a few enzymes to facilitate the use of a given substrate, in response to that substrate, the use of light as a sub&rate by dark-grown Euglena cells involves the coordinated induction of many enzyme activities and much structure, all properly coordinated, a process we call development. Substrate induction in microorganisms involves the recognition of substrate molecules by regulatory molecules which control transcription of the enzymes to be induced. It is entirely possible that pigmented regulatory molecules have also evolved which recognize specific wavelengths of light as inducing “substrates” (as in the control of photoreactivating en-

rRNA

215

TRANSCRIPTION

zymes (29)) and which control the transcription of molecules needed for plastid development, etc., such as the pl- and cytrRNAs. If this were so, these pigmented regulatory molecules would have evolved so that their absorptions came to resemble the absorptions of the molecules formed through their regulation. Thus it is possible that pigmented regulatory molecules have evolved to control the transcription of pl-rRNA whose absorption spectra are similar to or identical with the absorption of protochlorophyll(ide1, the normal chlorophyll precursor. Protochlorophyll(ide), absorbing as it does in the blue and the red regions of the spectrum, provides an absorption similar to chlorophyll, the molecule whose synthesis is controlled. This would make evolutionary sense since the control molecules should turn on chlorophyll synthesis or chloroplast development only when wavelengths of light are available which will permit photosynthesis once the photosynthetic apparatus is induced to form, if the control system is to be adaptive. At present we do not know which of the two models presented is correct or whether there are others. It does seem likely, however, that, if pigmented control molecules exist, we must expect a diversity of them with absorption spectra which resemble the pigments of the processes they control. REFERENCES 1. COHEN,

D., AND

SCHIFF,

J. A. (1973)Biophys.

J.

13, llla. 2. GREENBLATT, C. L., AND SCHIFF, J. A. (1959) J. Protozool. 6, 23-28. 3. SCHIFF, J. A. (1972) in Methods in Enzymology, Vol. 24B (San Pietro, A., ed.), Academic Press, New York. 4. COHEN, C., AND SCHIFF, J. A. (1974)PlantPhysiol. Suppl., 4. 5. STERN, A. I., SCHIFF, J. A., AND EPSTEIN, H. T. (1964) Plant Physiol. 39, 220-226. 6. RAWSON, J. R., AND STUTZ, E. (1969) Biochim. Biophys. Acta 190, 368-380. 7. HEIZMANN, P. (1970) Biochim. Biophys. Acta 224, 144-154. 8. RAWSON, J. R., AND STUTZ, E. (1968) J. Mol. Biol. 33, 309-314. 9. NOLL, H. (1969) Anal. Biochern. 27, 130-149. 10. PEACOCK, D. C., AND DINGMAN, C. W. (1968) Biochemistry 7, 668-932.

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11. ZELDIN, M. H., AND SCHIFF, J. A. (1967) Plant Physiol. 42, 922-932. 12. STERN, A. I., SCHIFF, J. A., AND EPSTEIN, H. T. (1965) Plant Physiol. 39, 226-231. 13. RAWSON, J. R., CROUSE, E. J., AND STUTZ, E. (1971) Biochim. Biophys. Actu 246, 507-516. 14. BROWN, R. D., AND HASELKORN, R. (1971) Proc. Nat. Acad. Sci. USA 68, 2536-2539. 15. COMB, D. G., AND ZEHAVI-VILNER, J. (1967) J. Mol. Biol. 28, 441-458. 16. PENE, J. J., KNIGHT, E., AND DARNELL, J. E. (1968) J. Mol. Biol. 33, 609-623. 17. DYER, T. A., AND LEECH, R. M. (1968) Biochem. J. 106, 689-698. 18. SCHIFF, J. A. (1974) in Proceedings. 3rd International Congress on Photosynthesis, Rehovoth (Avron, M., ed.), Vol. III, pp. 1691-1717. 19. SCOIT, N. S., GRAHAM, J., SMILLIE, R. M., AND MUNNS, R. (1971) in Autonomy and Biogenesis of Mitochondria and Chloroplasts (Boardman, M., ed.), p. 382, North-Holland, Amsterdam. 20. BOVARNICK, J. G., SCHIFF, J. A., FREEDMAN, Z., AND EGAN, J. M., JR. (1974) J. Gen. Microbial. 83, 63-71. 21. BOVARNICK, J. G., ZELDIN, M. H., AND SCHIFF, J. A. (1969) Develop. Biol. 19, 321-340. 22. SMILLIE, R. M., AND SCOTT, N. S. (1969) in Progress in Molecular and Subcellular Biology (Hahn, F. E., et al., eds.), Vol. 1, pp. 136-202, Springer-Verlag, New York. 23. LOEB, J. R., HOWELL, R. R., AND TOMKINS, G. M. (1965) Science 153, 531-534. 24. WEBER, M. (1972) Nature New Biol. 235, 58-60. 25. COCUCCI, S. M., AND SUSSMAN, M. (197O)J. Cell Biol. 45, 399-407. 26. FREYSSINET, G. P., HEIZMANN, G., TRABUCHET, G., AND NIGON, V. (1972) Physiol. Veg. 10, 421-428. 27. EGAN, J. M., JR., DORSKY, D., AND SCHIFF, J. A. (1975) Plant Physiol. 56, 318-323. 28. GOINS, D. J., REYNOLDS, R. J., SCHIFF, J. A., AND BARNETT, E. W. (1973) Proc. Nat. Acad. Sci. USA 70, 1749-1752. 29. DIAMOND, J., SCHIFF, J. A., AND KELNER, A. (1974) Arch. Biochem. Biophys. 167, 603-614.

SCHIFF 30. SCHWARTZBACH, S. D., SCHIFF, J. A., AND GOLDSTEIN, N. H. (1975) Plant Physiol. 56,313-317. 31. VAISBERG, A. J., SCHIFF, J. A., LI, L. AND FREEDMAN, Z. (1974) Plant Physiol. 57, 594601. 32. HEIZMANN, P. (1975) in Proceedings. 3rd International Congress on Photosynthesis, Rehovoth (Avron, M., ed.), Vol. III, pp. 1745-1754. 33. DARNELL, J. E. (1968) Bacterial. Rev. 32, 262290. 34. KAEMPFER, R. (1971) Proc. Nat. Acad. Sci. USA 68, 2458-2462. 35. HOLOWINSKY, A. W., AND SCHIFF, Plant Physiol. 45, 339-347. 36. ZELDIN, M. H., AND SCHIFF, 81, 1-15.

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40. NICHOLS, K. E., AND BOGORAD, L. (1962) Bot. Gaz. 124, 85. 41. BISHOP, N., AND SENGER, H. (1975) Presented at meetings on chloroplast development, University of Marburg, Marburg, Germany, 1975. 42. EDELMAN, (1965) J. 43. EGAN, J. Physiol. 44. SMILLIE, (1963)

M., SCHIFF, J. A., AND EPSTEIN, H. T. Mol. Biol. 11, 769-774. M., AND CARELL, E. F. (1972) Plant 50, 391-395.

R. M., EVANS, W. R., AND LYMAN, Brookhaven Symp. Biol. 16, 89.

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45. SCHWARTZBACH, S. D., FREYSSINET, G., AND SCHIFF, J. A. 0974) Plant Physiol. 53, 533542. 46. CROUSE, E., VANDREY, J., AND STUTZ, E. (1974) in Proceedings. 3rd International Congress on Photosynthesis, Rehovoth (Avron, M., ed.), Vol. III, pp. 1775-1786. 47. SCOTT, N. S. (1973) J. Mol. Biol. 81, 327-336. 48. TAMBURO, K. M., AND HUTNER, S. H. (1971) J. Protozool. 18, 667-671.