Comparative in vivo studies of RNA synthesis in the fetal liver, placenta, and maternal liver

Comparative in vivo studies of RNA synthesis in the fetal liver, placenta, and maternal liver T. TERRY HAYASHI DONG H. SHIN SAMANTHA WIAND Pittsburgh, Pennsylvania A study of simultaneous labeling of fetal liver, placenta, and maternal liver RNA was conducted using 20 day pregnant rats. After injection of a labeled precursor, the RNA was extracted by cold phenol technique and analyzed by sucrose gradient centrifugation. HC-ora.tic acid was incorporated readily into the RNA of all three tissues with the highest radioactivity in the fetal liver RNA followed by the placental RNA; 14C-uridine, however, was incorporated only into the fetal liver and the placental RNA. The labeled RNA isolated from the placentas of 20 day pregnant rats 12 hours after injection of either 14C-orotic acid or 14C-uridine showed an unusually reduced or absent JBS radioactivity peak associated with a normal absorbancy pattern indicating a preferential turnover of the 18S placental RNA synthesized during late gestation.

A L T H o u G H many facets of RNA metabolism have been investigated extensively in adult mammalian tissues, 2 • 11 • 12 • 17 there is still a paucity of information with regard to a similar or a simultaneous in vivo study of RNA metabolism of the fetus and the placenta. This has been mainly due to the technical difficulty of adequately labeling the fetal and placental RNA. Significant labeling of the fetal RNA with a specific precursor has been achieved only when the precursor is in~ jected directly into the fetal liver area 4 or into the amniotic sac.1 Such procedures have obvious disadvantages of using anesthetized animals and of injecting material into relatively small undefined areas. Previous works in this laboratory have From the Department of Obstetrics and Gynecology, University of Pittsburgh School of Medicine, Magee-Womens Hospital. This project was supported by Grant No. HD 01890 from the National Institutes of Health, United States Public Health Service, and was partly supported by Grant No. FR 05570 from the National Institutes of Health, United States Public Health Service.

1261

demonstrated that a variety of labeled precursors of RNA is transferred readily into the fetus of the near-term pregnant rat. 8 Additional studies9 have now indicated that the transplacental passage and the incorporation of the labeled precursors into RNA are influenced by the type of RNA precursors, the incubation time, and the duration of pregnancy. The RNA of the maternal liver, the fetus, and the placenta is labeled readily with orotic acid, but uridine is preferentially incorporated into the RNA of the fetus and the placenta. With both compounds, however, the incorporation of the precursors into the RNA did not take place until after the nineteenth day of pregnancy. The present investigation was undertaken using this prior information of optimum time of placental transfer and of active RNA incorporation to label the fetal and placental RNA with uc-<>rotic acid and 14C-uridine. A detailed analysis of such labeled RNA may afford a comparison of RNA from different age groups, i.e., the fetal liver RNA representing a young active embryonic tissue, the maternal liver RNA denoting a mature

1262 Hayashi, Shin, and Wiand

adult RNA, and the near-term placental RNA specifying senescent or old RNA.

Method Fifty to 100 p.c of a labeled precursor acid or l40-uridine, Schwarz BioResearch) was injected intraperitoneally into 20 day pregnant rats, Sprague~Dawley strain. At specified intervals the animals were killed by cervical dislocation. Maternal liver, fetal liver, and placentas were immediately removed and placed separately into 10 volumes of cold O.lM Na acetate buffer, pH 5.0, containing 0.5 per cent sodium dodecyl sulfate (SDS) and 0.1 per cent polyvinyl sulfate, and 10 volumes of water-saturated phenol.13 The preparations were homogenized in a Virtis Homogenizer at top speed for 20 seconds and placed on a Thomas Rotator for 1 hour. After centrifugation at 10,000 x g for 10 minutes, the aqueous, the interphase and the phenol layers were separated. To the aqueous layer ~ volume of phenol was added; the mixture was shaken in a Thomas Rotator for an additional ~ hour, and centrifuged at 10,000 x g for 10 minutes. Two and one-half volumes of 95 per cent ethanol and 7{ 0 volume of l.OM K acetate, pH 5.0, were added to the resulting aqueous layer, and RNA was precipitated overnight in a freezer. This preparation was labeled aqueous RNA and represents cytoplasmic RNAY To the interphase and the phenol layers, was added ~ volume of H 20 that was adjusted to 0.2 per cent with respect to SDS and 0.01 per cent with respect to hydroxyquinoline, para~aminosalicylic acid, and naphthalene disulfate. The mixture was shaken in a Burrell Shaker for 35 minutes and centrifuged at 10,000 x g for 10 minutes. One-half volume of phenol and ~ volume of water were added to the aqueous phase and the interphase. The mixture was shaken on a Burrell Shaker for 35 minutes. Again centrifugation was carried out at 10,000 x g for 10 minutes; 2.5 volume of 95 per cent ethanol and ~Ito volume of l.OM K acetate, pH 5, were added to the aqueous phase and RNA was allowed to precipitate overnight. This ( 14C-orotic

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December 15, 1969 J. Ohst. & Gynec.

preparation was labeled interphase RNA and represents primarily nuclear RNA. ~ The RNA suspensions from the aqueous and from the interphase extractions were centrifuged at 15,000 x g for 15 minutes to obtain RNA pellets; RNA ·was resuspended in 5 to 10 mi. of cold H~O and centrifuged at 10,000 x g for 10 minutes to remove any insoluble residues. The suspensions were made 0.02M with respect to MgCl 2 and 0.015M with respect to Tris HCI, pH 8.0, and then DNase (0.1 mg./1 ml. of RNA suspension) was added. The tubes were left in ice for one hour followed by addition of 2.5 volume of 95 per cent ethanol and 7{ 0 volume of l.OM K acetate, pH 5.0. The RNA was allowed to precipitate for 2 hours in a freezer and then centrifuged at 15,000 x g for 15 minutes. The resulting RNA pellets were resuspended in 2.0 mi. of H 2 0 and reprecipitated overnight with 2.5 volumes of 95 per cent ethanol and %0 volume of 1.0 M K acetate, pH 5.0. After centrifugation at 15,000 x g for 15 minutes the resulting RNA pellet was redissolved in 1.0 ml. of cold water. The absorbance at 260 mp. was determined, and approximately 0.5 to 1.0 mg. of RNA (absorbance at 260 mp. x 40 = p.g of RNA) 19 was placed on a 5 to 20 per cent sucrose gradient in l.OM K acetate, pH 5.0, as described previously.16 Centrifugation was conducted at 24,000 r.p.m. for 12 hours in a Spinco S.W. 25.1 rotor; gradients were cut into fractions of 1.0 ml. each after puncturing the bottom of the centrifuge tube with a needle. The absorbance at 260 mp. was measured in cold. To each 1.0 mi. fraction, 0.15 mg. of bovine serum albumin and 0.1 ml. of 50 per cent trichloroacetic acid were added; the resulting precipitate was then collected on a Millipore filter. The precipitate was washed 3 times with 3 ml. of cold 5 per cent trichloroacetic acid and placed on a planchet. The radioactivity was determined with a thin window gas flow detector. 1

Results Orotic acid. Fig. 1 represents the pattern of cytoplasmic RNAs obtained in a 20 day pregnant rat 12 hours after injection of 14 C-

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orotic acid. There is a high degree of labeling of the fetal liver RNA followed by moderate labeling of the placental and the maternal liver RNA. The radioactivity pattern corresponds well with the absorbance pattern, except for the placental RNA in which the specific activity of the 18S peak is much lower than that of the 28S peak. All the sucrose gradient analyses reported herein were repeated 4 to 5 times and in each instance similar radioactivity and absorbance patterns were obtained. The results of interphase or nuclear RNA extraction after orotic acid injection are presented in Fig. 2. The greatest radioactivity is again found in the fetal liver RNA, but its pattern is more heterogeneous indicating the presence of a messenger-like RNA. The other tissues show fairly well resolved radioactivity peaks that correspond rather closely with the absorbance pattern. It is not known how much of the radioactivity of the interphase RNA is due to contaminating cytoplas-

mic RNA. However, the contaminating RNA does not appear to interfere grossly with the radioactivity pattern of the nuclear RNA. The radioactivity pattern of the placental nuclear RNA, in contrast to the placental cytoplasmic RNA, displays the 18S radioactivity peak closely following the absorbance peak; therefore, the specific activity of the 18S peak does not differ from that of the 28S peak. The analysis of RNA after 30 and 60 minutes of pulse-labeling time displayed moderate radioactivity only in the interphase preparation. The radioactivity pattern displayed heterogeneous pattern suggesting messengerlike RNA formation as show in Fig. 3. Uridine. The pattern of labeled RNA after various pulse-labeling time with 14 C-uridine was similar to those observed after 14C-orotic acid except there was no significant radioactivity in the maternal liver RNA throughout the entire series of uridine experiments. The sucrose gradient centrifugation pat-

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clos, and Gonzalesl. with 140-orotic acid probably resulted from the use of 16 to 18 day pregnant rats. Likewise, the absence of any incorporation into the fetal RNA with 14C-adenine5 must have been due to the administration of the labeled precursor during the first half of pregnancy. Only a nonspecific precursor, such as 32 P, has been reported to be transferred across the placental barrier and be readily incorporated into the RNA of the fetal liver during early pregnancy.18 In the 12 hour experiments with 140orotic acid and 14 0-uridine, the highest incorporation was observed in the fetal liver RNA as one might anticipate since the fetal liver represents the most active tissue studied in our present work. In the uridine series, however, there was no significant labeling of the maternal liver. Additional studies with young male and young nonpregnant female rats weighing from 100 to 250 grams displayed significant labeling with 14C-labeled

terns of the cytoplasmic RNA of the fetal liver and placenta after a labeling time of 12 hours are seen in Fig. 4. The cytoplasmic RNA of the fetal liver displays an increase in specific activity with the radioactivity pattern closely following the absorbance pattern. Although the cytoplasmic RNA of the placenta shows increased total radioactivity, an unusual loss of 18S radioactivity is again observed. Fig. 5 shows the sucrose gradient separation of the cytoplasmic RNA after a relatively short labeling time of 60 minutes. Here there are definite and well defined 28S and 18S radioactivity peaks in both the placenta and the fetus. Comment

Our results clearly indicate that fetal liver and placental RNA can be labeled with a specific precursor if studies are performed on 20 day pregnant rats. The very limited degree of fetal labeling reported by Bresnick, Lan-

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1268 Hayashi, Shin, and Wiand

uridine. The pregnant animals used in the present work wen~ mated at 100 days of age and weighed between 300 to 400 grams. Studies are in progress to determine when the animal loses its ability to utilize uridine as a specific precursor for Jive1· RNA. A surprising observation in the prolonged labeling studies with 14 C~orotic acid and 14 0-uridine is the absence or the reduction of lBS radioactivity peak associated with a normal 18S absorbance pattern in the cytoplasmic RNA of the placenta. Since there is a definite 18S radioactivity peak after labeling time of 60 minutes with 14 0-uridine and since the 188 absorbance peak displays a normal pattern, the 18S ribosomal RNA synthesized in the placenta during a late gestation period may indicate a selective susceptibility to degradation. The formation of 288 and 188 RNA from 458 RNA in the nucleoli of HeLa cells has been studied in detail.l:;, 21 Present evidence indicates that in both the HeLa cells14 • 20 and in the rat liver10 the 18S RNA leaves the nucleus ahead of the 285 RNA. However,

REFERENCES

1. Bresnick, E., Lanclos, K., and Gonzales, E.: Biochim. Biophys. Acta 108: 568, 1965. 2. Bucher, N. L. R., and Swaffield, N. M.: Biochim. Biophys. Acta 108: 551, 1965. 3. Chaudhuri, S., and Lieberman, 1.: ]. Bioi. Chern. 243: 29, 1968. 4. Church, R., and McCarthy, B.: J. Molec. Bioi. 23: 477, 1967. 5. Dancis, J., and Balis, M. E.: J. Bioi. Chern. 207: 367, 1954. 6. Dessev, G. N., Markov, G. G., and Tsanev, R. G.: Life Sc. 5: 2331, 1966. 7. Ennis, H. L.: Molec. Pharmacal. 2: 543, 1966. B. Hayashi, T. T., and Garvey, B.: AM. ]. 0BsT. & GYNEC. 102: 1154, 1968. 9. Hayashi, T. T., Shin, D. H., and Wiand, S.: AM. J. 0BsT. & GvNEc. 102: 1144, 1968. 10. Henshaw, E. C., Revel, M. and Hiatt, H. H.: J. Molec. Biol. 14:: 241, 1965. I L Hiatt, H. H.: J. Molec. Bio1. 5: 217, 1962.

December 15, 1969 :\m. J. Ohst. & Gynec.

there is no report to explain the preferentia [ degradation of 18S RNA comparable to what we have observed in our placental cytoplasmic RNA after a prolonged labeling tinw. Henshaw, Revel, and Hiatr1° have found a marked detf•rioration of rapidly labeled 18S RNA, in contrast to stable ribosomal RNA, when extracting cytoplasmic RNA from wt liver; Dessev, Markov, and TsanevG have also reported similar selective degradation of DNA-like RNA in rat livN microsomes involving both 28 and 188 peaks. The works of Ennis' and Ohadhuri and Lieberman~ may be more closely related to our problem. They observed a decrease in 188 radioactivity peak in L cells and rat liver, respectively, after administration of cycloheximide. The unusual degradation of newly formed 18S ribosomal RNA peak may be related to protein synthesis activity; however, additional work is needed to determine whether the decrease in 18S radioactivity peak does indeed represent senescense and is characteristic of the aging process.

12. Kidson, C., and Kirby, K. S.: J. Molec. Bioi. 10: 187, 1964. 13. Kidson, C., Kirby, K. S., and Ralph, R. K.: J. Molec. Bioi. 7: 312, 1963. 14. Penman, S.: J. Malec. Biol. 17: 117, 1966. 15. Scherrer, K., Latham, H., and Darnell, J. F.: Proc. Nat. Acad. Sc. 49: 240, 1963. 16. Shin, D. H., and Moldave, K.: J. Molec. Bioi. 21: 231, 1966. 17. Siekevitz, P., Maggio, R., and Catalano, C.: Biochim. Biophys. Acta 129: 145, 1966. 18. Smellie, R. M. S., Mcindoe, W. M., Logan, R., Davidson, J. N., and Dawson, I. M.: Biochem. J. 54: 280, 1953. 19. Strauss, J., and Sinsheimer, R: J. Molec. Bioi. 7: 43, 1963. 20. Vaughn, M. H., Warner, J. R., and Darnell, J. E.: J. Molec. Bioi. 25: 235, 1967. 21. Warner, J., Soeiro, R., Birnboim, H. C., Girard, M., and Darnell, J. E.: J. M()Iec. Bioi. 19: 349, 1966.