Estimation of rRNA synthesis and degradation rates in senescing wheat leaves

ARCHIVES OF BIOCHEMISTRY AND BIOPHYSICS Vol. 260, No. 1, January, pp. 285-292,1988

Estimation

of t-RNA Synthesis and Degradation in Senescing Wheat Leaves’

Rates

LORENZO LAMATTINA, MARCELA PINEDO, V. PAULINE YUDI, RAFAEL F. PONT LEZICA,’ AND RUBEN D. CONDE In&it&o

de Investigaciones Biotigicas, Universidad National de Mar de1 Plata, and Centro de Investigaciones Biolbgicas, FIBA, C. Correo 1348, (7600) Mar de1 Plata, Argentina

Changes in cytoplasmic and chloroplast rRNA content and rates of rRNA synthesis and degradation of detached wheat leaves were determined. It was found that rRNA loss is proportionally higher in chloroplasts than in cytoplasm. Rates of synthesis were measured by incorporation of large amounts of [3H]orotic acid into rRNA. This approach overcame size differences between pyrimidine pools of cells under different physiological status. Furthermore, these pools reached nearly the same specific radioactivity as that of the administered solution. Rates of degradation were estimated either as the difference between synthesis and net variation of rRNA or by disappearance of radioactivity from 32P-labeled rRNA. Results indicated a decrease in the net rRNA synthesis capacity of leaves after 48 h of detachment. However, the fractional rates of rRNA synthesis were maintained in both cytoplasm and chloroplasts. Ribosoma1 RNA degradation rates were 2.5fold higher in chloroplast than in cytoplasm. The observed chloroplast rRNA loss is due to an increased degradation rate which is l&fold higher than the synthesis rate 48 h after detachment. o 1988Academicpress,I”~.

During senescence of both intact and excised leaves ribosomes are lost (l-5). This loss implies an rRNA loss and occurs at a higher rate in chloroplast rRNA than in cytoplasmic rRNA (4,6). Possible mechanisms involved in this process are either a decreased rate of rRNA synthesis and/ or an increased rate of rRNA degradation (7). Precise measurements of synthesis and degradation rates of rRNA from leaves have not been reported. However, the information available allows us to con-

sider the regulation of rRNA loss during senescence in terms of arrested synthesis and enhanced degradation (7,8). We have studied these possibilities in cytoplasmic and chloroplast rRNA of detached wheat leaves kept in darkness. Consequently, the rates of synthesis were estimated by labeling the rRNA after absorption of a large dose (nontrace amount) of [3H]orotic acid. The rates of rRNA breakdown were evaluated either by the difference between the rate of synthesis and the net rRNA variation or by examining the loss of radioactivity on rRNA previously labeled with [32P]orthophosphate.

i Supported by grants from the Consejo National de Investigaciones Cientificas y T&micas (CONICET) and the Comision de Investigaciones Cientificas de la Provincia de Buenos Aires (CIC), Argentina. This work is part of a Ph.D dissertation written by Lorenzo Lamattina, to be presented at the Universidad National de Mar del Plata, Argentina. ’ To whom correspondence should be addressed at present address: Department of Biology, Washington University, Campus Box 1137, St. Louis, MO 63130.

MATERIALS

AND

METHODS

Chemicals. [5-3H]Orotic acid (20 Ci/mmol) was from New England Nuclear (Boston, MA). [32P]Orthophosphoric acid in 0.02 HCl (carrier-free) was from the Comision National de Energia Atomica, Argentina. All others reagents came from Fluka (Buchs, Switzerland). 285

0003-9861/88

$3.00

Copyright 0 1988by AcademicPress,Inc. All rights of reproductionin any form reserved.

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Plant material. Wheat (Tm’ticum aestivum L. cv. San Agustin INTA) was generously supplied by the Balcarce Experimental Station of the Institute Nacional de Tecnologia Agropecuaria (INTA) Argentina. Plants were grown in vermiculite soaked in Hoagland nutrient solution at 25°C and under a 14-h photoperiod of 525 W/m2 irradiance supplied by Sylvania Gro-Lux fluorescent tubes. Thirteen days after sowing, the first leaves were fully expanded and had an average weight of 80 mg. At this moment they were detached and put in the dark at 25°C in tubes containing 0.5 ml of water. They were kept under these conditions for the times stated in each experiment. Water was changed every day. Preparation of total rRNA containing fraction. Five grams of leaves (about 60 leaves) was homogenized with 4 vol of a medium containing 100 mM KCl, 1 mM Mg acetate, and 100 mM Tris-HCl, pH 7.8, at 4°C in a Virtis homogenizer operated for 15 min at speed 20. This operation was performed in 5-min steps separated by 1-min resting periods. Then Triton X-100 was added to the homogenate at a final concentration of 3% (v/v). The suspension was centrifuged at 27,000g for 20 min. The precipitate was discarded. One milliliter of the supernatant was taken to determine the total leaf RNA content. The remaining was centrifuged in an MSE 60 Ti rotor at 100,OOOgfor 150 min. The supernatant was discarded. The precipitate was mixed with 20 vol of acetone at -20°C and left overnight. The precipitate was collected by centrifugation at 10,OOOgfor 10 min, resuspended in 1 ml of water, and clarified by centrifugation at 4000g for 15 min. The resulting supernatant was analyzed as rRNA preparation using 0.05-ml aliquots for the determination of either RNA mass or radioactivity. Determinaticm of RNA. Determinations were performed after alkaline hydrolysis of RNA as described by Fleck and Munro (9). Labeling solutions. [32P]Orthophosphate was dissolved in water to give a radioactivity of 0.7 &i/ml. When used in trace amounts, [3H]orotic acid was dissolved in water to give a radioactivity of 0.3 pCi/ml and adjusted to pH 7.0. When used in large amounts, [aH]orotic acid was dissolved in water with enough nonradioactive erotic acid to give a specific radioactivity of 0.4 &i/pmol (2 &i/ml) and adjusted to pH 7.0. Administration of radioactive precursors to leaves. Two leaves were put under light (525 W/m”) in tubes containing 0.1 ml of labeling solution and submitted to a strong air stream for 30 min. Thereafter, the air supply was stopped and the leaves remained under light for at least 90 min. Measurement of radioactivity incorporated into leaf rRNA. For both radioactivity measurement and estimation of rRNA content samples containing rRNA from 5 g of leaves were mixed with enough perchloric acid to obtain a 0.5 M concentration in 2 ml volume.

ET AL. After 1 h at 4”C, the samples were centrifuged at 2000g for 10 min. The supernatant was discarded. The precipitate was washed with 2 ml of 0.2 M perchloric acid. The pellets were mixed with 2 ml of 0.3 M KOH and incubated at 40°C for 1 h. Then, 2 ml of 0.8 M perchloric acid was added. After standing for 1 h at 4”C, they were centrifuged at 2,000g for 10 min. Two milliliters of the supernatant were mixed with 6 ml of a scintillation cocktail containing 33% (v/v) Triton X-100 and counted by scintillation spectrometry. The remaining supernatant was used either for the estimation of rRNA content or for the chromatographic analysis of the free nucleotides. Paper chromatographic separation of nucleotides. The chromatography was performed according to Wyatt (10). For this purpose, 1 ml of RNA hydrolysate was neutralized and concentrated to 0.05 ml with a nitrogen stream. The concentrate was spotted on Whatman No. 1 paper strips with AMP, UMP, GMP, and CMP as standards. Descending chromatography was performed with isopropanol:hydrochloric acid:water (170:41:39; by vol) as solvent. Samples and standard spots were located under ultraviolet light. They were cut and eluted with 1.2 ml of 0.1 N HCI, their absorbance was estimated at 260 nm, and they were mixed with 5 ml of a scintillation cocktail for counting. Isolation of ribosomal particles. The isolation of ribosomal particles from both chloroplast and cytoplasm was performed as indicated by Stutz and No11 (11). Twenty grams of leaves was homogenized with 3 vol of 50 mM KCI, 5 mM MgClz, 5 mM P-mercaptoethanol, 0.7 M sucrose, and 100 mM Tris-HCl, pH 7.5, at 4°C in a Waring blender homogenizer operated at its maximal speed for 15 s. This operation was performed in three 5-s periods separated by 1-min intervals. The homogenate was filtered through four cheesecloth layers and centrifuged at 400g for 2 min; the precipitate was discarded. The supernatant was centrifuged at 1lOOg for 12 min. The precipitate constitutes the chloroplast preparation. The supernatant was centrifuged at 26,000g for 30 min; the precipitate was discarded. The supernatant received Triton X-100 to a final concentration of 0.5% (v/v) and was layered on 2 ml of a medium containing 50 mM KCI, 5 mM MgClz, 5 mM P-mercaptoethanol, 1M sucrose, and 10 mM Tris-HCI, pH 7.5, and 4°C. Then it was centrifuged in a MSE 60 Ti rotor at 150,OOOg for 150 min. The precipitate was analyzed as total cytoplasmic ribosome preparation, The chloroplast preparation was resuspended in 8 ml of 5% (v/v) Triton X-100, 50 mM KCl, 5 mM MgClc, 5 mM p-mercaptoethanol, and 10 mM Tris-HCl at 4°C and centrifuged at 26,000g for 30 min; the precipitate was discarded. The supernatant was processed to obtain total chloroplast ribosomes by the same procedure described above for cytoplasmic ribosomes. Sucrose gradients. Cytoplasmic and chloroplast

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LEAF TABLE

rRNA

287

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I

INCORPORATION OF[3H]O~~~~~ACID INTOTOTAL LEAF rRNA THROUGHDIFFERENTLABELING APPROACHES rRNA synthesis

Expt.

Time after detachment (h)

rRNA (dpm * leaf ml. h-i)

incorporation (dpm . mg rRNA-’

. h-l)

1

0 24 48

51’7 f 65 3200 f 415 3093 f 505

2,651 +- 310 206,451 3~ 15,200 247,440 + 31,315

2

0 24 48

18.1 f 2.2 17.3 * 1.9 10.5 i 1.8

940 + 100 1,207 + 151 840 + 140

Net (ng * leaf -I. day-‘)

681 f 185 653 f 170 397 + 100

Fractional ( % . day-‘)

3.5 4.2 3.2

erotic Note. Sixty laves for each time after detachment were put under light to absorb 5 mM nonradioactive acid for 30 min and transferred to a 15 nM [3HJorotic acid (20 Ci/mmol) for labeling of RNA for 120 min (Expt. 1) or 5 mM [3H]orotic acid (0.4 Ci/mol) for 120 min (Expt. 2). The radioactivity incorporated per hour into rRNA was determined. Data were expressed either as dpm per leaf or as dpm per milligram of rRNA. The net rate of rRNA synthesis (Expt. 2) was calculated from the radioactivity incorporated, the specific radioactivity of the labeling solution, and the proportion of UMP in rRNA. Fractional rates are the percentages of total rRNA synthesized per day. Each value is the mean _+ SD of three experiments.

rRNA were analyzed by centrifugation in exponential sucrose gradients as in Stutz and No11 (11). For this purpose, cytoplasmic crude ribosomes were resuspended in 2 ml of 4 mM MgClz, 0.5% (w/v) SDS,3 10 mM Tris-HCl, pH 7.5, at 4°C and incubated at 37°C for 1 min. After SDS treatment, suspensions were layered on 10 ml of an exponential gradient of 0.3-l M sucrose in 4 mM MgClz, 1OmM Tris-HCl, pH 7.5, at 10°C. The gradients were centrifuged in a MSE SW Ti rotor at 150,OOOgfor 14.5 h according to Stutz and No11 (ll), and then scanned at 260 nm. In all cases, the centrifugal field stated in multiples of g is based on the average radius of rotation of the solution. RESULTS

Changes in total RNA and rRNA content.

Total leaf rRNA was measured at different times after detachment and the following values were obtained: zero time, 19.5 + 0.4 pg; first day, 16.3 + 0.5 pg and second day, 12.1 +- 1.0 pg. Hence, 2 days after detachment total rRNA was 62% of the initial value. This amounts to an rRNA average loss of 3.7 gg per day. In addition, rRNA comprises between 63 and 6’7% of total RNA in leaves at any time after detachment, suggesting that both total RNA and rRNA decrease at a similar rate. Di3 Abbreviation

used: SDS, sodium dodecyl sulfate.

rect measurements of total RNA and rRNA at different times confirmed this fact (data not shown). Incorporation of [‘Hlorotic acid into rRNA. The observed leaf RNA loss can be

attributed either to a decreased synthesis and/or to an increased degradation. Synthesis could be estimated by measuring the incorporation of trace amounts of labeled erotic acid into leaf rRNA. The disadvantage of the use of precursor in trace amounts rests on the fact that its ability to label RNA depends on changes of the pyrimidine pools as a consequence of the physiological status of cells (7, 12). We performed experiments using trace amounts of [3H]orotic acid in which rRNA label increases with time after detachment (not shown). Results in Table I show that this also occurs if leaves are absorbing large amounts of nonradioactive erotic acid during the 30 min previous to the administration of [3H]orotic acid in trace amounts. Hence, reduction of size differences between pyrimidine precursor pools of leaves at different times after detachment seems to depend on the constant absorption of large amounts of erotic acid. It has been described that when erotic acid is administrated in large amounts it is able

288

LAMATTINA

to expand the UTP pool in liver and kidney and thus the incorporation of labeled erotic acid into RNA became proportional to the synthetic capacity of the cells (13, 14). Hence, it seems possible to narrow size differences between pyrimidine precursor pools of plant cells by supplying to the leaves large amounts of labeled erotic acid and then estimating their capacity for rRNA synthesis. Under these conditions, it was found that incorporation of [3H]orotic acid into rRNA is linear for at least 3 h (data not shown). Then radioactivity incorporated into rRNA was determined after 2 h of labeling. Aliquots of hydrolyzed rRNA were also assayed chromatographically for determining both UMP proportion and radioactivity; the values obtained were 17 f 0.8 and 73 + 7%) respectively. The remaining 27% of the radioactivity was found in CMP. From those results the net rate of synthesis (ng leaf-’ * day-‘) was calculated as follows: the 73% of the dpm incorporated per leaf/ per day was divided by the specific activity of the labeling solution to obtain the nanomoles of UMP incorporated. It may be noted that, at different times after detachment, the amount of erotic acid absorbed per leaf during 2 h of labeling was zero time, 0.25 pmol; 2 days, 0.18 pmol. These amounts are on average 75-fold higher than the free pyrimidine nucleotides content in wheat leaves and in other plants (12). Hence, to estimate the amounts of rRNA synthesized, it is valid to assume that the specific activity of the precursor pool is the same as that of the labeling solution. Therefore, the nanomoles of UMP incorporated were multiplied by the molecular weight of UMP and divided by the fractional proportion of UMP in rRNA. The fractional rate of rRNA synthesis was calculated as the proportion existing between the net rRNA synthesis rate and the leaf rRNA content. Results in Table I show that when labeling was performed through a concentrated solution of [3H]orotic acid, the leaves retained their capacity for net rRNA synthesis during the first day after detachment. After that, this capacity decreases. Indirect estimation of total T-RNA degra-

ET AL.

dation rates. The synthesis values obtained in Table I were averaged for a l-day period and tabulated with the net changes in total rRNA content. Then the rates of rRNA degradation were estimated as the difference between synthesis and net rRNA variation. The results indicate that degradation is mainly responsible for the rRNA loss observed (Table II). This degradation rate increased 6-fold in the first 24 h over the synthesis rate. Identity of the lost rRNA. The observed rRNA loss could be either a selective process which preferentially affects only one rRNA type, or a process which affects similarly both cytoplasmic and chloroplastic rRNA. To investigate this point, a subcellular fractionation was performed from leaves at 0 to 48 h after detachment. The results in Fig. 1 indicate that rRNA loss was higher in chloroplast than in cytoplasm. This was confirmed by an additional experiment which shows the profiles of extracted rRNA from cytoplasm and chloroplast after centrifugation in exponential sucrose gradients. It indicates that 48 h after detachment rRNA loss is higher in chloroplast than in cytoplasm (Fig. 2). It also shows a heterogeneity in the distribution of light chloroplastic

TABLE

II

ESTIMATION OF FRACTIONAL RATES OF TOTAL rRNA DEGRADATION AS THE DIFFERENCE BETWEEN SYNTHESIS AND rRNA CONTENT VARIATION Time interval after detachment (h) O-24 24-48

Fractional rRNA synthesis 3.85 3.7

rates (% . day-‘) rRNA change -21.1 -26.4

rRNA degradation 24.95 30.1

Note. The fractional rates of synthesis values obtained in Table I at the initiation and at the end of each day were averaged. The net total rRNA changes were calculated as follows: the variation of rRNA over a l-day period was divided by the value of rRNA content in the middle of that period. Fractional rates of total rRNA degradation were calculated from the difference between synthesis and net rRNA changes.

Z’riticum

aestivum

LEAF

rRNA

TURNOVER

289

tic rRNA in a direct way (Table IV). Thus, degradation was followed by measuring the loss of radioactivity from rRNA previously labeled with [32P]orthophosphate. Results indicate that chloroplastic rRNA is degraded at a higher rate than cytoplasmic rRNA. DISCUSSION

Ribosomal RNA loss in detached wheat leaves represents an important fraction which amounts to 40% of the initial value 48 h after detachment. Similar results have been reported during senescence in several plant species (l-5,8,15,16). On the other hand, in contrast to protein and I

1 0 days

after

I

I

1

2

detachment 25 S

FIG. 1. Time course of rRNA changes in cytoplasm and chloroplast. Leaves were cut and put in darkness. At different times after detachment they were processed for determining both their cytoplasmic rRNA (0) and chloroplastic rRNA (0) contents as indicated under Materials and Methods. Zero time (100%) values were 11.9 pg of cytoplasmic rRNA per leaf and 3.4 pg of chloroplastic rRNA per leaf. Each value is the mean f SD of results obtained for two experiments

rRNA which could be due to a cleavage of 23 S rRNA (8). For that reason, 23 S rRNA loss appears disproportionate as compared to 16 S rRNA loss. Estimation of rRNA synthesis and degradation in cytoplasm and chloroplast. The

incorporation of radioactivity into cytoplasmic and chloroplastic rRNA was measured (Table III). The results indicate that absolute rates of synthesis decreased mainly in chloroplast. However, the fractional rates of synthesis did not change significantly. This indicates an increased degradation which is higher in chloroplastic rRNA than in cytoplasmic rRNA (2.5-fold). Direct estimation of rRNA degradation in cytoplasm and chloroplast. The above

estimations of rRNA degradation rates are indirect. Therefore, an experiment was performed to estimate the degradation rates of both cytoplasmic and chloroplas-

top

bottom

2. Sucrose gradient analysis of cytoplasmic FIG. and chloroplastic rRNA. Ribosomal RNA was extracted from cytoplasm (A) and chloroplast (B) of leaves obtained either at zero time (-) or at 48 h after detachment (---). Then they were analyzed by centrifugation in exponential sucrose gradients as described under Materials and Methods. Gradients were disassembled from the top and their profiles at 260 nm were recorded continuously.

LAMATTINA TABLE

ET AL. III

ESTIMATION OF RATES OF SYNTHESIS AND DEGRADATION OF CYTOPLASMIC AND CHLOROPLASTIC rRNA

Fraction

Net rRNA synthesis (ng * leaf ml* day-i)

Time after detachment (h)

Fractional Synthesis

rates of rRNA Change

(% * day-‘) Degradation

Cytoplasm

0 24 48

475 f 6 361 f 45 337 f 52

3.65 3.5

-11 -16

14.65 19.5

Chloroplast

0 24 48

90 f 19 91 f 20 37+ 5

3.3 3.35

-34 -50

37.3 53.35

Note. Leaves were labeled with large amounts of [3H]orotic acid as indicated in experiment 2 of Table I. They were processed to determine the radioactivity incorporated into eytoplasmic and chloroplastic rRNA. The net and fractional rates of synthesis were calculated as described in Table I. Degradation rates of rRNA were estimated as described in Table II. Each value is the mean k SD of results obtained for two experiments.

RNA loss, we have previously observed that DNA content remains constant up to 48 h after detachment in wheat leaves (17). Furthermore, during this period, protein loss could be retarded by treatment of the TABLE

IV

DECAY OF RADIOACTIVITY FROM rRNA LABELED WITH 32P

Fraction

Time after detachment (h)

[32P]rRNA (dpm . leaf -‘)

Loss of radioactivity (So)

Cytoplasm

5 48

5550 k 383 5193 + 433

0 6

Chloroplast

5 48

1667 t 240 947 + 80

0 43

Note. Leaves were labeled under light with [“Plorthophosphate for 120 min as indicated under Materials and Methods. Then they were washed and transferred to tubes containing 0.1 ml of 100 mM nonradioactive orthophosphate. Five hours after detachment, some of them were processed for estimation of radioactivity incorporated into cytoplasmic and chloroplastic rRNA. The remaining leaves were washed, transferred to tubes containing water, and put under darkness; 48 h after detachment they were processed for estimation of radioactive content in cytoplasmic and chloroplastic rRNA. Each value is the mean f SD of results obtained for two experiments.

leaves with kinetin (18). It seems that leaves retain their ability for the formation of complete ribosomal particles during the 48 h after detachment. Those previous studies suggested that our subsequent experiments should be performed during the 48 h after detachment. We looked first at the total leaf rRNA synthesis capacity. Synthesis is meant to include the production of mature rRNA molecules integrated to completed ribosoma1 particles. These experiments do not differentiate between effects on transcription and processing. To evaluate synthesis we measured the labeling of rRNA by [3H]orotic acid administration to the leaves. This precursor has been earlier used to label plant RNA (2). We found that the administration of trace amounts of [3H]orotic acid to the leaves was unsuitable for rRNA synthesis measurements. It gave paradoxical results with respect to the physiological status of leaves, probably due to a reduction in the size of intracellular pyrimidine pool with time after detachment (12). In contrast the labeling of rRNA during the constant uptake by the leaves of a concentrated solution of [3H]orotic acid allows us to obtain more consistent results than those obtained by using the same precursor in trace amounts. The apparent 6-fold increase in incorporation obtained by using the pre-

Triticum

aestivum

LEAF

cursor in trace amounts at 24 and 48 h after detachment was abolished by using the precursor in large amounts. Labeled erotic acid has been previously used in large amounts to evaluate the rRNA synthesis ability of animal cells (13,14). These large amounts abolish size differences between pyrimidine precursor pools of cells under different physiological conditions. Therefore, under these experimental conditions the data of incorporation of label into rRNA are independent of such size differences and should reflect the real intrinsic ability of the tissue to synthesize rRNA. The present report constitutes the first time such an approach has been used in plant tissues. Furthermore, we have converted also the incorporation of label data into mass of rRNA synthesized per leaf. The calculations are based on the assumption that the intracellular specific radioactivity of UTP is similar to that of the concentrated [3H]orotic acid solution administered. This assumption seems to be valid since the leaf pyrimidine nucleotide pool is enlarged Z-fold. In addition, we found that the fractional rates of rRNA synthesis did not change significantly between zero and 48 h after detachment. Hence, rRNA loss must be attributed to an increased rate of breakdown. Neither precise measurements of rates of turnover of the RNA fractions in leaves (19) nor reliable data on the rate of rRNA breakdown in detached leaves (7) have been provided. The exception is the work of Trewavas on Lemna minor (8) in which in vivo RNA turnover rates were measured by a different approach than that reported in the present work. They reported that when Lemna was incubated in water, rRNA was lost, as a consequence of both a reduction in the rate of synthesis and an increase in the rate of degradation. We report for the first time estimations of total rRNA degradation rates during induced senescence in wheat leaf and also the degradation process with regard to the type of degraded rRNA. We have thus concluded that the rate of rRNA breakdown is 2.5-fold higher in chloroplast than in cytoplasm.

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291

Either by indirect estimations of rRNA breakdown (Table III) or measurements of radioactivity loss from 32P-labeled rRNA (Table IV) data indicated that chloroplastic rRNA is degraded at a higher rate than cytoplasmic rRNA. However, it should be pointed out that direct estimations of degradation indicated that radioactivity in rRNA decayed at a lower rate than rRNA content in both cytoplasm and chloroplast. This is probably due to a recycling of the label which becomes a potentially serious problem during RNA studies (20, 21). On the other hand, indirect estimations of degradation are not influenced by recycling. Hence, in spite of errors arising from measurements of either rRNA synthesis or rRNA content, they provide more consistent results, and should be considered as a valid and useful method. It is thus concluded that loss of chloroplastic rRNA is mainly due to an increased breakdown. The results in Table III also indicate that fractional rates of synthesis are similar for both cytoplasmic and chloroplastic rRNA. If these rates are near to those of attached leaves, both kinds of rRNA would have similar in viva half-lives (about 19 days). This situation could be different from that reported by Trewavas for Lemna in which cytoplasmic rRNA and chloroplastic rRNA showed half-lives of 4 and 15 days, respectively (8). We have previously reported that the absolute rate of protein synthesis in leaves decreases after detachment (17). This decrease could be due partially to the chloroplastic rRNA loss observed. Brady and Steele-Scott (22) have reported a decreased capacity of the senescent wheat leaves of the synthesis of the large subunit of RuBPCase protein. They conclude that this occurred because chloroplastic ribosomes are more actively synthesizing other proteins than the large subunit of RuBPCase protein. Although the loss of chloroplastic rRNA probably reflects an increased breakdown of complete ribosomes, this point needs to be investigated further by measuring the rate of breakdown of chloroplast ribosoma1 protein.

LAMATTINA ACKNOWLEDGMENTS We are indebted to the members of the Instituto de Investigaciones Biologicas, with a special mention to August0 Garcia, for helpful discussions and encouragement. Special thanks are also given to Barbara Schaal and Joseph E. Varner, Department of Biology, Washington University, for critical reading of the manuscript. L.L. holds a postgraduate scholarship from the Consejo National de Investigaciones Cientificas y TQcnicas (CONICET); R.P.L. and R.D.C. are established researchers from the Comisidn de Investigaciones Cientificas de la Prov. de Buenos Aires (CIC) and CONICET, respectively.

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12. WEINSTEIN,L. H., MCCUNE,D. C., MANCINI, J. F., AND VAN LEUKEN, P. (1969) Plant Physiol. 44, 1499-1510.

13. BUCHER,M. L. R., AND SWAFFIELD,M. N. (1969) Biochim Biophys. Acta 174,491-502. 14. IAPALUCCI-ESPINOZA,S., BUR,J. A., PUCCIARELLI, M. G., AND CONDE,R. D. (1986) Amer. J Physiol. 251, E266-272. 15. WOLLGIEHN,R. (1967) Symp. Sot. Exp. Biol. 21, REFERENCES 231-246. 16. WOLLGIEHN, R., LERBS, S., AND MUNSCHE, D. 1. WOLLGIEHN,R. (1961) Flora 151,411-437. (1976) Biochem. Physiol. PjZanzen. 170, 2. OSBORNE,D. J. (1962) Plant Physiol. 37,595-602. 3. EILAM, Y., BUTLER, R. D., AND SIMON, E. W. 381-387. (1971) Plant Physiol. 47,31’7-323. 17. LAMATTINA, L., PONT LEZICA, R. F., AND CONDE, R. D. (1985) Plant Physiol. 77,587-590. 4. TAKEGAMI, T. (1975) Plant Cell Physiol. 16, 18. LAMATTINA, L., ANCHOVERRI,V., CONDE,R. D., 417-425. AND PONT LEZICA, R. F. (1987) Plant Physiol. 5. NAITO, K., IIDA, A., SUZUKI, H., AND TSUI, H. (1979) Physiol. Plant 46,50-53. 83,497-499. 6. CALLOW,J. A., CALLOW,M. E., AND WOOLHOUSE, 19. WOOLHOUSE,H. W. (1982) in The Molecular BiolH. W. (1972) Cell oi$ 1, 79-90. ogy of Plant Development (Smith, H., and Grierson, D., Eds.), pp. 256-281, Blackwell, 7. FARKAS, G. L. (1982) in Encyclopedia of Plant Physiology: Nucleic Acids and Proteins in Oxford. Plants II (Parthier, B., and Boulter, D., Eds.), 20. WATTS,J. W., AND HARRIS, H. (1959) B&hem, J. New Series, Vol. 14B, pp. 224-262, Springer72,147-153. 21. MANDELSTAM,J. (1963) Ann. N. Y. Acad. Sci. 102, Verlag, Berlin/Heidelberg/New York. 8. TREWAVAS,A. (1970) Plant Physiol. 45,742-751. 61. 9. FLECK, A., AND MUNRO, H. N. (1962) B&him. 22. BRADY, C. J., AND STEELE-SCOTT,N. (1977) Aust. Biophys. Acta 55, 571-583. J. Plant PhysioL 4,327-335.