- Email: [email protected]
DNA polymerases in midgestation mouse embryo, trophoblast, and decidua
DEVELOPMENTAL
BIOLOGY
34, 200-210 (1973)
DNA Polymerases
in Midgestation
Mouse
Embryo,
Trophoblast,
and Decidua MICHAEL I. SHERMAN AND HYEN SAM KANG’ Department
of Cell Biology,
Roche Institute Accepted
of Molecular April
Biology,
Nutley,
New Jersey 07110
20, 1973
The DNA polymerases of midgestation mouse embryo, trophoblast, and decidua have been examined. A low molecular weight, nuclear, DNA-dependent polymerase (D-DNA polymerase) and a higher molecular weight cytoplasmic enzyme were found in all three cell types. A DNA polymerase which utilized the poly(A) strand of oligo(dT) .poly(A) as template (R-DNA polymerase) was also found in the three cell types. This enzyme was present both in the nucleus and the cytoplasm. All enzyme levels were highest in the rapidly dividing embryonic cells, substantially lower in the DNA replicating but nondividing trophoblast cells, and lowest in the nonreplicating, nondividing decidual cells. Our observations are consistent with the idea that the nuclear and cytoplasmic D-DNA polymerases are under coordinate control. The relationship of these enzymes to DNA synthesis in uivo is discussed. INTRODUCTION
The cells of the early mammalian embryo divide rapidly and have a relatively short DNA synthetic (S) phase (Graham, 1973). On the other hand, giant trophoblast cells, which are of embryonic origin but are located in the placenta, do not undergo cell division in rodents (Alden, 1948; Zybina, 1960; Atlas et al., 1960; Zavarzin et al., 1966). The nuclei of these cells accumulate large amounts of DNA in the first half of gestation (Zybina, 1961, 1964; Hunt and Avery, 1971; Barlow and Sherman, 1972). This phenomenon is due neither to engulfment of maternal cells (Barlow and Sherman, 1972) nor to fusion of trophoblast cells (Chapman et al., 1972; Gearhart and Mintz, 1972). At present, the most likely explanation for the formation of polyploid trophoblast nuclei is endoreduplication, i.e., cycles of replication of the entire genome in the absence of mitosis (Zybina and Mos’yan, 1967; Sherman et al., 1972). The DNA polymerases in trophoblast, or other polyploid cells, have not been charac1Present address: The Wistar Institute, 36th Street and Spruce, Philadelphia, Pennsylvania 19104.
terized. We have undertaken a study of the DNA-dependent DNA polymerases (hereafter designated D-DNA polymerase) in 11th day mouse trophoblast cells. Recently, Temin (1971) proposed that RNA-dependent DNA polymerases might be implicated in cell differentiation. A number of investigators have now provided evidence for the presence of synthetic RNA-dependent DNA polymerase activity in chick and sea urchin embryos (Stavrianopoulos et al., 1971; Maia et al., 1971; Slater and Slater, 1972), and in adult mammalian tissues (Ward et al., 1972; Bolden et al., 1973). We have, therefore, included in our studies the characterization of an enzyme in embryonic and maternal cells which is capable of copying the poly(A) strand of oligo(dT) . poly(A). This enzyme will be referred to as R-DNA polymerase (Fridlender et al., 1972). The trophoblast enzymes were compared with those observed in the embryo proper and decidua at the same stage of pregnancy. The latter cells constitute the maternal moiety of the placenta and are nonreplicating and nondividing at this gestation age (Atlas et al., 1960; Sherman, unpublished observations). By this choice 200
Copyright All rights
0 1973 by Academic Press, Inc. of reproduction in any form reserved
SHERMAN AND KANC
Mouse Embryonic DNA Polymerases
of cells, we have been able to ask whether qualitative or quantitative enzyme differences exist between diploid embryo and polyploid trophoblast cells and between rapidly dividing embryonic and nondividing maternal cells. MATERIALS
AND
METHODS
Materials. Unlabeled deoxynucleotides, salmon sperm DNA, and dithiothreitol (DTT) were purchased from Sigma Chemicals, St. Louis, Missouri. Poly(A) and bovine serum albumin were obtained from Miles Laboratories, Kankakee, Illinois. Oligo(dT), 12-18 nucleotides in length, was purchased from Collaborative Research Inc., Waltham, Massachusetts. [3H]dTTP was purchased from New England Nuclear, Boston, Massachusetts. Preparation of tissues. Embryos were obtained 11.5 days after random mating of SWR/J p x SJL/J 8 mice (Jackson Laboratories, Bar Harbor, Maine). Embryo, trophoblast, and decidua were dissected apart in phosphate-buffered saline (solution A of Dulbecco; Gibco, New York) as described previously (Sherman, 1972). Yolk sacs were discarded. Pooled tissues from 20 to 100 conceptuses were washed and frozen in isotonic sucrose solution (1 mM KPO, pH 6.8, 2 mM MgCl, , 0.5 mM DTT, 0.32 M sucrose). After thawing, the cells were disrupted in a Dounce homogenizer. In order to minimize breakage of giant trophoblast nuclei in those experiments where separate nuclear and cytoplasmic fractions were required, the following procedure was adopted and used for all the tissues. The homogenate was centrifuged at 500 rpm for 4 min to pellet unbroken cells, debris, and some nuclei. This pellet was repeatedly resuspended, gently homogenized, and centrifuged at low speed until clumps of tissue were no longer visible by microscopic observation (usually 3-4 cycles). All the fractions were then pooled. Nuclei were pelleted and
201
washed by centrifugation at 1500 g for 10 min. The pooled supernatants constitute the cytoplasmic fraction. Mitochondria were removed from this fraction by centrifugation for 15 min at 25,000 g. The nuclear pellets were treated basically as described by Chang and Bollum (1971). Nuclei were disrupted by homogenization in 2 M NaCl followed by stirring for 2 hr at 4”. This step also extracted some of the DNA which was removed by centrifugation at 105,000 g for 4 hr. Both nuclear and cytoplasmic fractions were then dialyzed against hypotonic phosphate buffer (0.02 M KPO, pH 7.4, 0.5 mA4 DTT), and clarified by centrifugation at 15,000 g for 10 min. When fractionation of the cells was not required, NaCl to a final concentration of 2 M was added directly to the whole homogenate, and the procedure then followed was that described above for the nuclear fraction. DEAE-cellulose chromatography. DEAEcellulose columns (Whatman No. 52) preequilibrated with 0.02 M KPO, pH 7.4, 0.5 mM DTT were used for chromatographic separation of the enzymes. The samples usually contained about 20 mg protein, and the ratio of protein applied/ column size was 2-4 mg/ml packed cellulose. After adsorption of the protein, the columns were washed with 25 ml of KPO, buffer. Adhering protein was then eluted with a linear 0.02 to 0.5 M KPO, gradient at pH 7.4 containing 0.5 mM DTT. The gradient volume was 100 to 120 ml (ca. 10 column volumes). Fifty microliters of each fraction were assayed for enzyme activity. Prior to glycerol gradient centrifugation, the preparations were usually passed through a preparative DEAE-cellulose column. In this case, after high salt treatment and dialysis, the extracts were applied to the column in 0.02 M KPO, buffer, pH 7.4, and directly eluted with 0.4 M phosphate buffer. Under these conditions, polymerases eluted from the column, while at least
202
DEVELOPMENTAL BIOLOGY
some of the residual DNA was retained. After dialysis against 0.02 M phosphate buffer, the samples were prepared for centrifugation as described below. Glycerol density gradients. In order to concentrate the samples for gradient centrifugation after passage through DEAEcellulose columns, the nuclear, cytoplasmic, or whole cell extracts were treated with (NH,),SO, at 70% saturation (Chang and Bollum, 1971), followed by dialysis of the resuspended pellet against 0.02 M phosphate buffer. The samples were then layered on a 5 ml, 10-30s glycerol gradient containing 0.02 M KPO, pH 7.4, 0.5 mM DTT and 0.1% Triton X-100 (or sodium deoxycholate). Centrifugation was carried out at 45,000 rpm in a Spinco SW 5OL rotor at 4°C for 14 hr. Samples were collected from the bottom of the tube, and 50-~1 aliquots were assayed for enzyme activity. Sedimentation coefficients and molecular weights were estimated from the cosedimenting protein markers chymotrypsinogen, ribonuclease, ovalbumin, and aldolase (Pharmacia Fine Chemicals, Piscataway, New Jersey). Enzyme assays. Activated DNA used in the D-DNA polymerase assay was prepared according to Schlabach et al. (1971). The oligo(dT) . poly(A) template utilized in the R-DNA polymerase assay was prepared as described by Fridlender et al. (1972). D-DNA polymerase assays were carried out at 37” according to Schlabach et al. (1971). The following 200~~1reaction mixtures were used: 0.05 M Tris .HCl pH 8.5, 0.01 M MgClz, 0.5 mM DTT, 90 pg of bovine serum albumin, 100 pg of nicked, native salmon sperm DNA, 0.1 mM each dATP, dGTP, and dCTP, 0.025 mM [3H]dTTP, and 50 ~1 of the appropriate enzyme fraction. The specific activity of the [3H]dTTP was between 10 and 100 cpm/pmole, except when high specific activity substrate (15,000 cpm/pmole) was used to detect enzymes which might be
VOLUME 34, 1973
present at low levels or with low activities. The incubation period was 30-60 min. R-DNA polymerase assays were basically those used by Fry and Weissbach (1973) for the mouse L-cell enzyme. Assays were carried out at 30°C for 30 or 60 min using the following components in a total volume of 200 ~1: 0.05 M Tris .HCl pH 7.5, 0.125 M KCl, 0.5 mM MnCl,, 2.5 mM DTT, 90 pg of bovine serum albumin, 5 pg of oligo(dT) .poly(A), and 0.013 mM [3H]dTTP (specific activity 100-200 cpm/ pmole, except when high specific activity substrate was used). After incubation in the D-DNA polymerase assay, the tubes were placed in ice and 250 pg of heat denatured carrier salmon sperm DNA were added, followed by cold 5% trichloroacetic acid (TCA). The contents of each tube were filtered through TCA-washed glass fiber disks (Whatman GF/A, 2.4 cm). The disks were then washed three times with 5% TCA followed by two washes with 80% ethanol. After 15 min under an infrared lamp, the disks were counted by liquid scintillation spectrometry in Liquifluor solution (New England Nuclear). The preparation of samples for counting in the R-DNA polymerase assay was the same, except that after the incubation period, the tubes were placed in ice and a solution (0.3 ml) containing 0.056 M sodium pyrophosphate and 50 PM unlabeled dTTP as well as carrier DNA (250 pg) was added to each tube before TCA precipitation. Also, the disks were presoaked for 5 min in 0.1 M sodium pyrophosphate. These additions were found to considerably reduce the background in the R-DNA polymerase assay. RESULTS
Characterization
of D-DNA
Polymerases
When whole cell extracts of embryo, trophoblast, or decidua were analyzed by
SHERMAN AND KANC
Mouse Embryonic
DEAE-cellulose column chromatography, a similar profile was observed in all three cases: only one region of D-DNA polymerase activity was observed, eluting at about 0.14 A4 salt. A typical experiment is shown in Fig. 1. Split peaks or shoulders were observed in about one-third of the profiles and may reflect enzyme heterogeneity as noted previously by Chang and Bollum (1971). When nuclei and cytoplasm were separated and chromatographed individually, both gave profiles very similar to those of the whole cell extracts. No further
DNA Polymerases
203
peaks of enzyme activity were observed when very high specific activity substrate (ca. 15,000 cpm/pmole) was used. Chang and Bollum (1971) have demonstrated that extracts from a variety of mammalian cell types gave two peaks of D-DNA polymerase activity after centrifugation on sucrose gradients. When mouse embryonic tissues were prepared by their procedure and then centrifuged on glycerol gradients, almost all of the activity sedimented in a single peak. If, however, the preparations were first passed through a preparative DEAE-cellulose column in order to more thoroughly remove endogenous DNA (see Materials and Methods), and then centrifuged, two discrete peaks of activity were observed in the gradient in all three tissues (Fig. 2). The approximate sedimentation coefficients were 2.5-3 S and 6-7 S, corresponding to molecular weights of about 25,000 and 120,000, respectively. In these preparations, a small amount of activity was present between the two peaks. This activity may be associated with the R-DNA polymerase which sediments in this region of the gradient (see below). When nuclei and cytoplasm were analyzed separately, a striking difference in the profiles was observed (Fig. 3). While the cytoplasmic extract contained only rapidly sedimenting D-DNA polymerase activity, approximately 90% of the nuclear activity was present as the slow sedimenting species. The enzymes were similarly distributed in the three cell types studied.
Characterization of R-DNA Polymerase I
40
FRACTION
60 NUMBER
FIG. 1. DEAE-cellulose column chromatography of D-DNA polymerases. Whole cell extracts were prepared, adsorbed on a DEAE-cellulose column, and eluted with a linear, 0.02 to 0.5 M KPO, gradient. Fifty-microliter aliquots of each fraction (2.1 ml) were assayed for D-DNA polymerase activity. (a) embryo; (b) trophoblast; (c) decidua.
When homogenates were prepared for glycerol gradient analysis as in Fig. 2, the bulk of R-DNA polymerase activity in trophoblast, embryo, and decidua sedimented between the two D-DNA polymerase peaks (Fig. 4). The estimated sedimentation coefficient was 4-4.5 S, corresponding to a molecular weight of approximately
204
DEVELOPMENTAL BIOLOGY
VOLUME 34, 1973
FRACTION
NUMBER
FIG. 3. Glycerol density gradient profiles of nuclear and cytoplasmic D-DNA polymerases. Extracts were prepared from decidual tissue as described in Materials and Methods, passed through a preparative DEAE-cellulose column, centrifuged, and assayed for D-DNA polymerase activity. O--O, nuclear extract; e---O, cytoplasmic extract. 6
5 IO 15 20 T FRACTION NUMBER
FIG. 2. Glycerol density gradient profiles of DDNA polymerases from embryo, trophoblast, and decidua. Whole-cell extracts, after passage through a preparative DEAE-cellulose column, were treated as described in Materials and Methods. (a) embryo; (b) trophoblast; (c) decidua.
65,000. In all three cases, varying amounts of activity sedimented faster than the main peak. This may have been due to aggregation or adherence to residual DNA in the extracts (see Discussion). The R-DNA polymerase activity could also be separated from the bulk of D-DNA polymerase activity if the extracts were passed through a preparative DEAE-cellulose column to remove residual DNA and then chromatographed on a second DEAE-cellulose column using a salt gradient for elution (Fig. 5). Under these conditions, both the embryo and trophoblast R-DNA polymerase eluted at 0.08-0.09 M salt, whereas the D-DNA polymerase eluted at 0.14 M. No further peaks of R-DNA polymerase activity were observed when high specific activity substrate was used in the enzyme assay.
Template Polymerase
Specificity
of
the
R-DNA
The template specificity of R-DNA polymerase partially purified by chromatography on DEAE-cellulose is described in Table 1. The enzyme appeared to use poly(dT) . poly(A) more efficiently than oligo(dT) .poly(A). We have, however, routinely used the latter as a template in R-DNA polymerase assays because it cannot be utilized by the D-DNA polymerases, whereas the former template can be used, albeit poorly, by HeLa and L-cell D-DNA polymerases (Fridlender et al., 1972; Fry and Weissbach, 1973). Although the enzyme was unable to incorporate dTTP into acid-insoluble material when poly(A) . poly(U) or poly(A) alone was used as template, activity was observed when oligo(A) .poly(U) was added to the assay mix. The lack of incorporation of substrate in the presence of poly(A) or oligo(dT) alone argues against the possibility that the enzyme is a terminal transferase. Many of the properties of the R-DNA polymerase de-
SHERMAN AND KANG
Mouse Embryonic
DNA Polymerases
205
FRACTION NUMBER FIG. 4. Glycerol density gradient profiles of RDNA polymerase. Whole cell extracts were treated as described in Fig. 2 and then assayed for R-DNA polymerase activity (see Materials and Methods). The arrows indicate the location of the high molecular weight (fraction 7) and low molecular weight (fraction 19) D-DNA polymerase peak fractions. (a) trophoblast; (b) embryo; (c) decidua.
scribed here are similar to those observed in other embryonic cell types (Maia et al., 1971; Stavrianopoulos et al., 1971; Slater and Slater, 1972). Fry and Weissbach (1973) have found that the L-cell R-DNA polymerase is also able to utilize an oligo(dG) -poly(C) template, about 10% as efficiently as oligo(dT) .poly(A). Furthermore, activated, native DNA was about half as efficient as oligo(dT) .poly(A). Our results with oligo(dG) poly(C) were variable, but in most cases, the R-DNA polymerase studied here was able to use this template about 5-15% as efficiently as oligo(dT) .poly(A). We are unable to determine with certainty the extent to which the R-DNA polymerase can utilize an activated, native DNA template because of possible contamination of our R-DNA polymerase preparations with D-DNA polymerases.
FRACTIONNUMBER FIG. 5. DEAE-cellulose column chromatography of R-DNA polymerases. Whole cell extracts were treated as described in Fig. 1 except that they were first passed through a preparative DEAE-cellulose column. The R-DNA polymerase assay procedure is described in Materials and Methods. The arrows refer to the position of the peak tube of D-DNA polymerase activity. (a) embryo; (b) trophoblast.
Quantitation
of
Polymerase Activities
In order to obtain a meaningful measure of DNA polymerase levels in our samples, it was necessary to eliminate nuclease activity. When crude homogenates were assayed, labeled dTTP was incorporated into acid-insoluble material. However, the amount of incorporation rose and then dropped with increasing incubation times or amounts of homogenate. This was not observed in assays with partially purified enzyme. Enzyme activity was linear for at least 2 hr following glycerol gradient centrifugation, and for at least 40 min following DEAE-cellulose chromatography. In the latter case, although the activity leveled off, the labeled product was not reduced, even after 2 hr (Fig. 6). Furthermore, labeled single-stranded or double-
206
DEVELOPMENTAL BIOLOGY
VOLUME 34, 1973
stranded DNA added to the standard assay mix was only very slightly degraded, if at all, to acid soluble material after two hours of incubation. Although the glycerol gradient enzymes exhibited linear kinetics for longer periods of time than the enzymes eluted from DEAE-cellulose, activity was consistently 6-10 times higher in the latter preparations. Consequently, the enzymes partially purified by DEAE-cellulose chromatography were used for quantitative analyses.
Table 2 is a compilation of our results. Experiment 1 indicates that D-DNA polymerase activity was largely cytoplasmic, while the R-DNA polymerase was more evenly distributed between nucleus and cytoplasm. Both embryo and trophoblast had the same distributions. The specific activities of both D-DNA and R-DNA polymerases were higher in the embryo than in the trophoblast or decidua (experiment 2). To determine whether levels of DNA polymerase in trophoblast cells doubled TABLE 1 with every doubling of DNA during enTEMPLATE SPECIFICITY OF MOUSE R-DNA POLYMERASE doreduplication, the ratio of DNA polymerase activity per milligram DNA was Expt. Template Activity” 9% No. Activity calculated. This ratio was 7-9 times higher for the D-DNA polymerase and 4-6 times 1 Oligo(dT) .poly(rA) 770 100 higher for the R-DNA polymerase in the Poly(dT) .poly(rA) 1677 218 embryo than the trophoblast (Table 2). Oligo(rU) .poly(rA) 436 57 Poly( rU) poly( rA) Substantial differences were also found to 51 7 Poly(rA) 63 8 exist between the embryo and decidua DNA polymerase : DNA ratios. The specific 2 Oligo(dT) .poly(rA) 5658 100 activity differences between embryo, troPoly(dT) .poly(rA) 8535 151 phoblast, and decidua were larger when Oligo(dT) 39
Incubation
Time (Mid
FIG. 6. Kinetics of the D-DNA and R-DNA polymerase reactions in embryo and trophoblast. The preparations used had been chromatographed on DEAE-cellulose columns. Assay conditions are described in Materials and Methods. (a) D-DNA polymerase; (b) R-DNA polymerase. O-0, Embryo 04, trophoblast.
SHERMAN AND KANC
Mouse Embryonic TABLE
LEVELS OF D-DNA
AND R-DNA
Nuclear
Embryo Trophoblast Experiment 2 Embryo ~e;plmi;~last
Embryo Fe;I$$last
2
POLYMERASEACTIVITY IN ELEVENTH DAY EMBRYO, TROPHOBLAST,AND DECIDUA % Activity
Experiment 1 Embryo Trophoblast
207
DNA Polymerases
Specific activity” (pmoles/min/mg)
Cytoplasmic
Protein
DNA
Embryokrophoblast (or embryo/decidua) Protein*
DNA’
22 23
D-DNA 78 77
Polymerase -d -
-
2.9
7:
:z
R-DNA 56 54
Polymerase 2.4 1.4
71 16
1.7
4.5
D-DNA
Polymerase 170 52 19
4868 541 266
31 8.9
9.0 18.3
Polymerase 5.1 2.6 1.8
145 27 25
2; 2.8
-
R-DNA -
Et:
a Specific activities were calculated on the basis of the protein (Lowry et al., 1951) or DNA (Burton, 1956) content of the initial untreated homogenate. bThe ratio of specific enzyme activity on a per milligram protein basis in the embryo to that in trophoblast or decidua. c The ratio of specific enzyme activity on a per milligram DNA basis in the embryo to that in trophoblast or decidua. d Because high specific activity [3H]dTTP was used in this experiment, substrate concentrations were too low to provide a reliable estimate of the specific enzyme activity.
DISCUSSION
In the course of our experiments, we did not find any qualitative differences in the D-DNA and R-DNA polymerases in mouse embryo, trophoblast, or decidua on the 11th day of gestation. Such differences might be present at other gestation ages. It would, however, be difficult to so expand our studies because of the problems involved in separating trophoblast from decidua at later stages (Sherman, 1972) and obtaining adequate amounts of tissue at earlier stages. Although the nuclear and cytoplasmic D-DNA polymerases were separable under appropriate conditions by glycerol gradient centrifugation, they appeared to cochromatograph on DEAE-cellulose. In contrast, Weissbach et al. (1971) found two peaks of D-DNA polymerase activity when HeLa cell nuclei were chromatographed on
a DEAE-cellulose column: 1540% of the activity passed unadsorbed through the column in the 0.02 M phosphate buffer wash, and the remainder of the activity eluted at higher salt concentrations. Recently, using relatively large quantities of tissue, we observed in nuclear, but not in cytoplasmic, extracts, some D-DNA polymerase activity eluting in the wash fraction from a DEAE-cellulose column (following passage through an initial preparative column). However, even under these conditions, a substantial fraction of the activity was retained on the column. The results presented here suggest that in our hands the nuclear D-DNA and the R-DNA polymerase show a strong affinity for residual endogenous DNA; unless the DNA is removed, this interaction affects the chromatographic and sedimentation profiles of the enzymes. A further complication might
208
DEVELOPMENTAL BIOLOGY
be the possible interconversion of the high and low molecular weight D-DNA polymerases, suggested by the recent studies of Chang and Bollum (1972b) and Hecht (1972). It should, nevertheless, be noted that in any given experiment, regardless of the purification procedure, the sedimentation or chromatographic profiles were the same in the three tissues under study. Weissbach et al. (1971) reported that HeLa nuclear extracts contained both low and high molecular weight D-DNA polymerases while the cytoplasmic extract contained only the latter. Chang and Bollum (1972a), on the other hand, observed that rat liver nuclei contained only the low molecular weight enzyme, while the cytoplasmic extract contained both enzymes. Finally, Hecht (1972) found that in the mouse testis, nuclear extracts contained only the low molecular weight D-DNA polymerase, while the cytoplasmic extract contained only the high molecular weight species. Our results are in close agreement with those of Hecht (1972). The discrepancies may be due to the different species studied, or the method of preparing the nuclear and cytoplasmic extracts. Fry and Weissbach (1973) observed that the R-DNA polymerase of mouse L-cells in culture was largely cytoplasmic. As Table 2 indicates, we have found an equal distribution of the enzyme between nucleus and cytoplasm. The significance of this difference awaits further study. Autoradiographic studies indicate that the S phase in giant trophoblast cells is relatively short and that all giant cells are capable of incorporating 3H-thymidine into acid-insoluble material (Cameron, 1964; Zavarzin et al., 1966; Sherman, unpublished observations). The data in Table 2, however, show that the specific activities of R-DNA polymerase and total D-DNA polymerase in embryo were twice and three times as high, respectively, as in trophoblast. These differences were amplified when calculated on a per milligram of DNA basis (Table 2). The differences in specific
VOLUME34, 1973
activity were not due to cochromatography of inhibitors in the trophoblast preparation or activators in the embryo preparation, since activity was additive when the trophoblast and embryo enzymes were mixed and assayed (unpublished results). If the levels of the DNA polymerases were rising proportionately with the accumulation of DNA in trophoblast giant cells, the ratio of enzyme activity to DNA content should be similar to that observed in embryo cells. On the other hand, if there were no increase in the levels of DNA polymerase as polyploidization occurred, this ratio in the embryo preparation would be expected to be between 13 and 39 times that in the trophoblast preparation, calculated on the basis of 10-30s of the trophoblast cells having undergone polyploidization and reached ploidy levels averaging 256n by the 11th day of gestation (Barlow and Sherman, 1972). The intermediate values obtained for both D-DNA (7-9 times) and R-DNA (4-6 times) polymerases suggest that some increase in the enzyme levels had occurred during polyploidization, but not to the extent required to maintain the enzyme: DNA ratios. If the DNA polymerases studied here are largely responsible for replication of the genome, then relatively low levels of enzyme would appear to be adequate to replicate the greatly enlarged amount of DNA in trophoblast giant nuclei in a reasonably short time. It would then follow that the levels of DNA polymerase present in diploid mouse embryonic cells are far in excess of that required for replication. It must be pointed out that the relationship between the enzymes studied in this report, and in others, to replication in uiuo has yet to be established. Grippo and Lo Scavo (1972) have provided evidence that a new DNA polymerase activity appears in mature Xenopus laevis oocytes in preparation for the onset of replication. Chang and Bollum (1972a) have observed that only the 6-8 S cytoplasmic DNA polymerase activity rises significantly during rat liver
SHERMAN AND KANG
Mouse Embryonic
regeneration, leading them to propose that this enzyme is involved in DNA replication in c&o. Fansler and Loeb (1969) observed a redistribution of DNA polymerase from cytoplasm to nucleus during embryogenesis in the sea urchin. The cell types in the present study are either (a) replicating and dividing, (b) replicating but not dividing, or (c) neither replicating nor dividing. Yet we have not observed in any of these cases either the presence of a unique enzyme, any striking difference in the relative proportions of the enzymes, or any significant variation in the distribution of the enzymes between nucleus and cytoplasm. It would appear from our experiments that the nuclear and cytoplasmic D-DNA polymerases are coordinately controlled, both being present at high levels in actively dividing cells, and at lower levels in nondividing cells. The role of the R-DNA polymerase in in uiuo DNA synthesis is at present ambiguous. Fry and Weissbach (1973) have carried out a detailed study on an enzyme in mouse L cells with properties very similar to those described here. Although this enzyme is not a terminal transferase and can utilize a number of artificial templates (see also Table l), in vitro conditions have not yet been found which will enable the enzyme to use bacteriophage, viral, or natural mRNA as template (Fry and Weissbach, 1973). Nevertheless, none of our data refutes Temin’s hypothesis that an R-DNA polymerase, perhaps the enzyme described here, plays a role in cell differentiation (Temin, 1971). If this were the case, however, the presence of significant amounts of the same enzyme in decidua, a maternal, nondividing tissue would have to be explained. We wish to thank Ms. N. Chew, Dr. A. Weissbach, and especially Dr. M. Fry for their help. REFERENCES ALDEN, R. H. (1948). Implantation of the rat egg. III. Origin and development of primary trophoblast cells. Amer. J. Anat. 83. 143-182.
DNA Polymeruses
209
ATLAS,M., BOND, V. P., and CRONKITE, E. P. (1960). DNA synthesis in the developing mouse embryo studied with tritiated thymidine. J. Histochem. Cytochem. 8, 171-181. BARLOW, P. W., and SHERMAN, M. I. (1972). The biochemistry of differentiation of mouse trophoblast: studies on polyploidy. J. Embryol. Exp. Morphol. 27, 447-465. BOLDEN, A., FRY, M., MULLER, R., and WEISSBACH,A. (1972). The presence of a polyriboadenylic aciddependent DNA polymerase in eukaryotic cells. AFC~. Biochem. Biophys. 153, 26-33. BURTON, K. (1956). A study of the conditions and mechanism of the diphenylamine reaction for the calorimetric estimation of the deoxyribonucleic acid. Biochem. J. 62, 315-323. CAMERON, I. L. (1964). Is the duration of DNA synthesis in somatic cells of mammals and birds constant? J. Cell Biol. 20, 185-188. CHANG, L. M. S., and BOLLUM, F. J. (1971). Low molecular weight deoxyribonucleic acid polymerase in mammalian cells. J. Biol. Chem. 246,5835-5837. CHANG, L. M. S., and BOLLUM, F. J. (1972a). Variation of deoxyribonucleic acid polymerase activities during rat liver regeneration. J. Biol. Chem. 247, 7948-7950. CHANG, L. M. S., and BOLLUM, F. J. (197213). Antigenie relationships in mammalian DNA polymerase. Scence 175, 1116-1117. CHAPMAN, V. M., ANSELL, J. D., and MCLAREN, A. (1972). Trophoblast giant cell differentiation in the mouse: expression of glucose phosphate isomerase (GPI-1) electrophoretic variants in transferred and chimeric embryos. Deuelop. Biol. 29, 48-54. FANSLER, B., and LOEB, L. A. (1969). Sea urchin nuclear DNA polymerase. II. Changing localization during early development. Exp. Cell Res. 57, 305-310. FRIDLENDER, B., FRY, M., BOLDEN, A., and WEISSBACH, A. (1972). A new synthetic RNA-dependent DNA polymerase from human tissue culture cells. Proc. Nat. Acad. Sci. U.S. 69, 452-455. FRY, M., and WEISSBACH,A. (1973). The utilization of synthetic DNA templates by a new DNA polymerase from cultured murine cells. J. Biol. Chem. 248, 2678-2683. GEARHART, J. D., and MINTZ, B. (1972). Glucosephosphate isomerase subunit-reassociation tests for maternal-fetal and fetal-fetal cell fusion in the mouse placenta. Develop. Biol. 29, 55-64. GRAHAM, C. F. (1973). The cell cycle during mammalian development. In “The Cell Cycle in Development and Differentiation” (M. Balls and F. S. Billett, eds.), pp. 293-310. Cambridge Univ. Press, London and New York. In press. GRIPPO, P., and Lo SCAVO,A. (1972). DNA polymerase activity during maturation in Xenopus laevis oocytes. Biochem. Biophys. Res. Commun. 48, 280285.
210
DEVELOPMENTAL BIOLOGY
HECHT, N. B. (1972). The interconvertibility of mouse DNA polymerase activities derived from the nucleus and cytosol. J. Cell Biol. 55, 109a. HUNT, C. V., and AVERY, G. B. (1971). Increased levels of deoxyribonucleic acid during trophoblast giant cell formation in mice. J. &prod. pert. 25, 85-94. LOWRY, 0. H., ROSEBROUGH,N. J., FARR, A. L., and RANDALL, R. J. (1951). Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265-275. MAIA, J. C. C., ROUGEON, F., and CHAPEVILLE, F. (1971). Chick embryo poly(rA:dT)-dependent DNA polymerase. FEBS Letr. 18, 130-134. SCHLABACH, A., FRIDLENDE~ B., BOLDEN, A., and WEISSBACH, A. (1971). DNA-dependent DNA polymerases from HeLa cell nuclei. II. Template and substrate utilization. Biochem. Biophys. Res. Com-
mun. 44,879-885. SHERMAN, M. I. (1972). The biochemistry of differentiation of mouse tropboblast: alkaline phosphatase.
Develop. Biol. 27, 337-350. SHERMAN, M. I., WALKER, P. M. B., and MCLAREN, A. (1972). Mechanism of accumulation of DNA in Nature (London) giant cells of mouse trophoblast. New Biol. 238, 175-176. SLATER, I., and SLATER, D. W. (1972). DNA polymerase potentials of sea urchin embryos. Nature (London) New Biol. 27, 81-84. STAVRIANOPOULOS,J. G., KARKAS, J. D., and CHARGAFF,
VOLUME 34, 1973
E. (1971). Nucleic acid polymerases of the developing chicken embryo: a DNA polymerase preferring a hybrid template. Proc. Nat. Acad. Sci. U.S. 68, 2207-2211. TEMIN, H. M. (1971). The protovirus hypothesis. J. Nut. Can. Inst. 46: III-VII. WARD, D. C., HUMPHREYS, K. C., and WEINSTEIN, I. B. (1972) Synthetic RNA-dependent DNA polymerase activity in normal rat liver and hepatomas.
Nature (London) 237, 499-503. WEISSBACH, A., SCHLABACH, A., FRIDLENDER, B., and BOLDEN, A. (1971). DNA polymerases from human cells. Nature (London) New Biol. 231, 167-170. ZAVARZIN, A. A., SAMOSHKINA, N. A., and DONDUA, A. K. (1966) Synthesis of DNA and kinetics of cells in early embryogenesis of mice. Zh. Obshch. Biol. 27,
697-709. ZYBINA, E. V. (1960). Sex chromatin in the trophoblast of early white rat embryos. Dokl. Akad. Nauk.
SSSR 130, 633-635. ZYBINA, E. V. (1961). Endomitosis and polyteny of trophoblast giant cells. Dokl. Akad. Nauk. SSSR 140, 1177-1180. ZYBINA, E. V. (1963). Cytophotometric determination of polyploidization of the cells of the trophoblast. Dokl. Akad. Nauk. SSSR 153, 1428-1431. ZYBINA, E. V., and MOS’YAN, I. A. (1967). Sex chromatin bodies during endomitotic polyploidication of trophoblast cells. Tsitologiya 9, 265-272.
BIOLOGY
34, 200-210 (1973)
DNA Polymerases
in Midgestation
Mouse
Embryo,
Trophoblast,
and Decidua MICHAEL I. SHERMAN AND HYEN SAM KANG’ Department
of Cell Biology,
Roche Institute Accepted
of Molecular April
Biology,
Nutley,
New Jersey 07110
20, 1973
The DNA polymerases of midgestation mouse embryo, trophoblast, and decidua have been examined. A low molecular weight, nuclear, DNA-dependent polymerase (D-DNA polymerase) and a higher molecular weight cytoplasmic enzyme were found in all three cell types. A DNA polymerase which utilized the poly(A) strand of oligo(dT) .poly(A) as template (R-DNA polymerase) was also found in the three cell types. This enzyme was present both in the nucleus and the cytoplasm. All enzyme levels were highest in the rapidly dividing embryonic cells, substantially lower in the DNA replicating but nondividing trophoblast cells, and lowest in the nonreplicating, nondividing decidual cells. Our observations are consistent with the idea that the nuclear and cytoplasmic D-DNA polymerases are under coordinate control. The relationship of these enzymes to DNA synthesis in uivo is discussed. INTRODUCTION
The cells of the early mammalian embryo divide rapidly and have a relatively short DNA synthetic (S) phase (Graham, 1973). On the other hand, giant trophoblast cells, which are of embryonic origin but are located in the placenta, do not undergo cell division in rodents (Alden, 1948; Zybina, 1960; Atlas et al., 1960; Zavarzin et al., 1966). The nuclei of these cells accumulate large amounts of DNA in the first half of gestation (Zybina, 1961, 1964; Hunt and Avery, 1971; Barlow and Sherman, 1972). This phenomenon is due neither to engulfment of maternal cells (Barlow and Sherman, 1972) nor to fusion of trophoblast cells (Chapman et al., 1972; Gearhart and Mintz, 1972). At present, the most likely explanation for the formation of polyploid trophoblast nuclei is endoreduplication, i.e., cycles of replication of the entire genome in the absence of mitosis (Zybina and Mos’yan, 1967; Sherman et al., 1972). The DNA polymerases in trophoblast, or other polyploid cells, have not been charac1Present address: The Wistar Institute, 36th Street and Spruce, Philadelphia, Pennsylvania 19104.
terized. We have undertaken a study of the DNA-dependent DNA polymerases (hereafter designated D-DNA polymerase) in 11th day mouse trophoblast cells. Recently, Temin (1971) proposed that RNA-dependent DNA polymerases might be implicated in cell differentiation. A number of investigators have now provided evidence for the presence of synthetic RNA-dependent DNA polymerase activity in chick and sea urchin embryos (Stavrianopoulos et al., 1971; Maia et al., 1971; Slater and Slater, 1972), and in adult mammalian tissues (Ward et al., 1972; Bolden et al., 1973). We have, therefore, included in our studies the characterization of an enzyme in embryonic and maternal cells which is capable of copying the poly(A) strand of oligo(dT) . poly(A). This enzyme will be referred to as R-DNA polymerase (Fridlender et al., 1972). The trophoblast enzymes were compared with those observed in the embryo proper and decidua at the same stage of pregnancy. The latter cells constitute the maternal moiety of the placenta and are nonreplicating and nondividing at this gestation age (Atlas et al., 1960; Sherman, unpublished observations). By this choice 200
Copyright All rights
0 1973 by Academic Press, Inc. of reproduction in any form reserved
SHERMAN AND KANC
Mouse Embryonic DNA Polymerases
of cells, we have been able to ask whether qualitative or quantitative enzyme differences exist between diploid embryo and polyploid trophoblast cells and between rapidly dividing embryonic and nondividing maternal cells. MATERIALS
AND
METHODS
Materials. Unlabeled deoxynucleotides, salmon sperm DNA, and dithiothreitol (DTT) were purchased from Sigma Chemicals, St. Louis, Missouri. Poly(A) and bovine serum albumin were obtained from Miles Laboratories, Kankakee, Illinois. Oligo(dT), 12-18 nucleotides in length, was purchased from Collaborative Research Inc., Waltham, Massachusetts. [3H]dTTP was purchased from New England Nuclear, Boston, Massachusetts. Preparation of tissues. Embryos were obtained 11.5 days after random mating of SWR/J p x SJL/J 8 mice (Jackson Laboratories, Bar Harbor, Maine). Embryo, trophoblast, and decidua were dissected apart in phosphate-buffered saline (solution A of Dulbecco; Gibco, New York) as described previously (Sherman, 1972). Yolk sacs were discarded. Pooled tissues from 20 to 100 conceptuses were washed and frozen in isotonic sucrose solution (1 mM KPO, pH 6.8, 2 mM MgCl, , 0.5 mM DTT, 0.32 M sucrose). After thawing, the cells were disrupted in a Dounce homogenizer. In order to minimize breakage of giant trophoblast nuclei in those experiments where separate nuclear and cytoplasmic fractions were required, the following procedure was adopted and used for all the tissues. The homogenate was centrifuged at 500 rpm for 4 min to pellet unbroken cells, debris, and some nuclei. This pellet was repeatedly resuspended, gently homogenized, and centrifuged at low speed until clumps of tissue were no longer visible by microscopic observation (usually 3-4 cycles). All the fractions were then pooled. Nuclei were pelleted and
201
washed by centrifugation at 1500 g for 10 min. The pooled supernatants constitute the cytoplasmic fraction. Mitochondria were removed from this fraction by centrifugation for 15 min at 25,000 g. The nuclear pellets were treated basically as described by Chang and Bollum (1971). Nuclei were disrupted by homogenization in 2 M NaCl followed by stirring for 2 hr at 4”. This step also extracted some of the DNA which was removed by centrifugation at 105,000 g for 4 hr. Both nuclear and cytoplasmic fractions were then dialyzed against hypotonic phosphate buffer (0.02 M KPO, pH 7.4, 0.5 mA4 DTT), and clarified by centrifugation at 15,000 g for 10 min. When fractionation of the cells was not required, NaCl to a final concentration of 2 M was added directly to the whole homogenate, and the procedure then followed was that described above for the nuclear fraction. DEAE-cellulose chromatography. DEAEcellulose columns (Whatman No. 52) preequilibrated with 0.02 M KPO, pH 7.4, 0.5 mM DTT were used for chromatographic separation of the enzymes. The samples usually contained about 20 mg protein, and the ratio of protein applied/ column size was 2-4 mg/ml packed cellulose. After adsorption of the protein, the columns were washed with 25 ml of KPO, buffer. Adhering protein was then eluted with a linear 0.02 to 0.5 M KPO, gradient at pH 7.4 containing 0.5 mM DTT. The gradient volume was 100 to 120 ml (ca. 10 column volumes). Fifty microliters of each fraction were assayed for enzyme activity. Prior to glycerol gradient centrifugation, the preparations were usually passed through a preparative DEAE-cellulose column. In this case, after high salt treatment and dialysis, the extracts were applied to the column in 0.02 M KPO, buffer, pH 7.4, and directly eluted with 0.4 M phosphate buffer. Under these conditions, polymerases eluted from the column, while at least
202
DEVELOPMENTAL BIOLOGY
some of the residual DNA was retained. After dialysis against 0.02 M phosphate buffer, the samples were prepared for centrifugation as described below. Glycerol density gradients. In order to concentrate the samples for gradient centrifugation after passage through DEAEcellulose columns, the nuclear, cytoplasmic, or whole cell extracts were treated with (NH,),SO, at 70% saturation (Chang and Bollum, 1971), followed by dialysis of the resuspended pellet against 0.02 M phosphate buffer. The samples were then layered on a 5 ml, 10-30s glycerol gradient containing 0.02 M KPO, pH 7.4, 0.5 mM DTT and 0.1% Triton X-100 (or sodium deoxycholate). Centrifugation was carried out at 45,000 rpm in a Spinco SW 5OL rotor at 4°C for 14 hr. Samples were collected from the bottom of the tube, and 50-~1 aliquots were assayed for enzyme activity. Sedimentation coefficients and molecular weights were estimated from the cosedimenting protein markers chymotrypsinogen, ribonuclease, ovalbumin, and aldolase (Pharmacia Fine Chemicals, Piscataway, New Jersey). Enzyme assays. Activated DNA used in the D-DNA polymerase assay was prepared according to Schlabach et al. (1971). The oligo(dT) . poly(A) template utilized in the R-DNA polymerase assay was prepared as described by Fridlender et al. (1972). D-DNA polymerase assays were carried out at 37” according to Schlabach et al. (1971). The following 200~~1reaction mixtures were used: 0.05 M Tris .HCl pH 8.5, 0.01 M MgClz, 0.5 mM DTT, 90 pg of bovine serum albumin, 100 pg of nicked, native salmon sperm DNA, 0.1 mM each dATP, dGTP, and dCTP, 0.025 mM [3H]dTTP, and 50 ~1 of the appropriate enzyme fraction. The specific activity of the [3H]dTTP was between 10 and 100 cpm/pmole, except when high specific activity substrate (15,000 cpm/pmole) was used to detect enzymes which might be
VOLUME 34, 1973
present at low levels or with low activities. The incubation period was 30-60 min. R-DNA polymerase assays were basically those used by Fry and Weissbach (1973) for the mouse L-cell enzyme. Assays were carried out at 30°C for 30 or 60 min using the following components in a total volume of 200 ~1: 0.05 M Tris .HCl pH 7.5, 0.125 M KCl, 0.5 mM MnCl,, 2.5 mM DTT, 90 pg of bovine serum albumin, 5 pg of oligo(dT) .poly(A), and 0.013 mM [3H]dTTP (specific activity 100-200 cpm/ pmole, except when high specific activity substrate was used). After incubation in the D-DNA polymerase assay, the tubes were placed in ice and 250 pg of heat denatured carrier salmon sperm DNA were added, followed by cold 5% trichloroacetic acid (TCA). The contents of each tube were filtered through TCA-washed glass fiber disks (Whatman GF/A, 2.4 cm). The disks were then washed three times with 5% TCA followed by two washes with 80% ethanol. After 15 min under an infrared lamp, the disks were counted by liquid scintillation spectrometry in Liquifluor solution (New England Nuclear). The preparation of samples for counting in the R-DNA polymerase assay was the same, except that after the incubation period, the tubes were placed in ice and a solution (0.3 ml) containing 0.056 M sodium pyrophosphate and 50 PM unlabeled dTTP as well as carrier DNA (250 pg) was added to each tube before TCA precipitation. Also, the disks were presoaked for 5 min in 0.1 M sodium pyrophosphate. These additions were found to considerably reduce the background in the R-DNA polymerase assay. RESULTS
Characterization
of D-DNA
Polymerases
When whole cell extracts of embryo, trophoblast, or decidua were analyzed by
SHERMAN AND KANC
Mouse Embryonic
DEAE-cellulose column chromatography, a similar profile was observed in all three cases: only one region of D-DNA polymerase activity was observed, eluting at about 0.14 A4 salt. A typical experiment is shown in Fig. 1. Split peaks or shoulders were observed in about one-third of the profiles and may reflect enzyme heterogeneity as noted previously by Chang and Bollum (1971). When nuclei and cytoplasm were separated and chromatographed individually, both gave profiles very similar to those of the whole cell extracts. No further
DNA Polymerases
203
peaks of enzyme activity were observed when very high specific activity substrate (ca. 15,000 cpm/pmole) was used. Chang and Bollum (1971) have demonstrated that extracts from a variety of mammalian cell types gave two peaks of D-DNA polymerase activity after centrifugation on sucrose gradients. When mouse embryonic tissues were prepared by their procedure and then centrifuged on glycerol gradients, almost all of the activity sedimented in a single peak. If, however, the preparations were first passed through a preparative DEAE-cellulose column in order to more thoroughly remove endogenous DNA (see Materials and Methods), and then centrifuged, two discrete peaks of activity were observed in the gradient in all three tissues (Fig. 2). The approximate sedimentation coefficients were 2.5-3 S and 6-7 S, corresponding to molecular weights of about 25,000 and 120,000, respectively. In these preparations, a small amount of activity was present between the two peaks. This activity may be associated with the R-DNA polymerase which sediments in this region of the gradient (see below). When nuclei and cytoplasm were analyzed separately, a striking difference in the profiles was observed (Fig. 3). While the cytoplasmic extract contained only rapidly sedimenting D-DNA polymerase activity, approximately 90% of the nuclear activity was present as the slow sedimenting species. The enzymes were similarly distributed in the three cell types studied.
Characterization of R-DNA Polymerase I
40
FRACTION
60 NUMBER
FIG. 1. DEAE-cellulose column chromatography of D-DNA polymerases. Whole cell extracts were prepared, adsorbed on a DEAE-cellulose column, and eluted with a linear, 0.02 to 0.5 M KPO, gradient. Fifty-microliter aliquots of each fraction (2.1 ml) were assayed for D-DNA polymerase activity. (a) embryo; (b) trophoblast; (c) decidua.
When homogenates were prepared for glycerol gradient analysis as in Fig. 2, the bulk of R-DNA polymerase activity in trophoblast, embryo, and decidua sedimented between the two D-DNA polymerase peaks (Fig. 4). The estimated sedimentation coefficient was 4-4.5 S, corresponding to a molecular weight of approximately
204
DEVELOPMENTAL BIOLOGY
VOLUME 34, 1973
FRACTION
NUMBER
FIG. 3. Glycerol density gradient profiles of nuclear and cytoplasmic D-DNA polymerases. Extracts were prepared from decidual tissue as described in Materials and Methods, passed through a preparative DEAE-cellulose column, centrifuged, and assayed for D-DNA polymerase activity. O--O, nuclear extract; e---O, cytoplasmic extract. 6
5 IO 15 20 T FRACTION NUMBER
FIG. 2. Glycerol density gradient profiles of DDNA polymerases from embryo, trophoblast, and decidua. Whole-cell extracts, after passage through a preparative DEAE-cellulose column, were treated as described in Materials and Methods. (a) embryo; (b) trophoblast; (c) decidua.
65,000. In all three cases, varying amounts of activity sedimented faster than the main peak. This may have been due to aggregation or adherence to residual DNA in the extracts (see Discussion). The R-DNA polymerase activity could also be separated from the bulk of D-DNA polymerase activity if the extracts were passed through a preparative DEAE-cellulose column to remove residual DNA and then chromatographed on a second DEAE-cellulose column using a salt gradient for elution (Fig. 5). Under these conditions, both the embryo and trophoblast R-DNA polymerase eluted at 0.08-0.09 M salt, whereas the D-DNA polymerase eluted at 0.14 M. No further peaks of R-DNA polymerase activity were observed when high specific activity substrate was used in the enzyme assay.
Template Polymerase
Specificity
of
the
R-DNA
The template specificity of R-DNA polymerase partially purified by chromatography on DEAE-cellulose is described in Table 1. The enzyme appeared to use poly(dT) . poly(A) more efficiently than oligo(dT) .poly(A). We have, however, routinely used the latter as a template in R-DNA polymerase assays because it cannot be utilized by the D-DNA polymerases, whereas the former template can be used, albeit poorly, by HeLa and L-cell D-DNA polymerases (Fridlender et al., 1972; Fry and Weissbach, 1973). Although the enzyme was unable to incorporate dTTP into acid-insoluble material when poly(A) . poly(U) or poly(A) alone was used as template, activity was observed when oligo(A) .poly(U) was added to the assay mix. The lack of incorporation of substrate in the presence of poly(A) or oligo(dT) alone argues against the possibility that the enzyme is a terminal transferase. Many of the properties of the R-DNA polymerase de-
SHERMAN AND KANG
Mouse Embryonic
DNA Polymerases
205
FRACTION NUMBER FIG. 4. Glycerol density gradient profiles of RDNA polymerase. Whole cell extracts were treated as described in Fig. 2 and then assayed for R-DNA polymerase activity (see Materials and Methods). The arrows indicate the location of the high molecular weight (fraction 7) and low molecular weight (fraction 19) D-DNA polymerase peak fractions. (a) trophoblast; (b) embryo; (c) decidua.
scribed here are similar to those observed in other embryonic cell types (Maia et al., 1971; Stavrianopoulos et al., 1971; Slater and Slater, 1972). Fry and Weissbach (1973) have found that the L-cell R-DNA polymerase is also able to utilize an oligo(dG) -poly(C) template, about 10% as efficiently as oligo(dT) .poly(A). Furthermore, activated, native DNA was about half as efficient as oligo(dT) .poly(A). Our results with oligo(dG) poly(C) were variable, but in most cases, the R-DNA polymerase studied here was able to use this template about 5-15% as efficiently as oligo(dT) .poly(A). We are unable to determine with certainty the extent to which the R-DNA polymerase can utilize an activated, native DNA template because of possible contamination of our R-DNA polymerase preparations with D-DNA polymerases.
FRACTIONNUMBER FIG. 5. DEAE-cellulose column chromatography of R-DNA polymerases. Whole cell extracts were treated as described in Fig. 1 except that they were first passed through a preparative DEAE-cellulose column. The R-DNA polymerase assay procedure is described in Materials and Methods. The arrows refer to the position of the peak tube of D-DNA polymerase activity. (a) embryo; (b) trophoblast.
Quantitation
of
Polymerase Activities
In order to obtain a meaningful measure of DNA polymerase levels in our samples, it was necessary to eliminate nuclease activity. When crude homogenates were assayed, labeled dTTP was incorporated into acid-insoluble material. However, the amount of incorporation rose and then dropped with increasing incubation times or amounts of homogenate. This was not observed in assays with partially purified enzyme. Enzyme activity was linear for at least 2 hr following glycerol gradient centrifugation, and for at least 40 min following DEAE-cellulose chromatography. In the latter case, although the activity leveled off, the labeled product was not reduced, even after 2 hr (Fig. 6). Furthermore, labeled single-stranded or double-
206
DEVELOPMENTAL BIOLOGY
VOLUME 34, 1973
stranded DNA added to the standard assay mix was only very slightly degraded, if at all, to acid soluble material after two hours of incubation. Although the glycerol gradient enzymes exhibited linear kinetics for longer periods of time than the enzymes eluted from DEAE-cellulose, activity was consistently 6-10 times higher in the latter preparations. Consequently, the enzymes partially purified by DEAE-cellulose chromatography were used for quantitative analyses.
Table 2 is a compilation of our results. Experiment 1 indicates that D-DNA polymerase activity was largely cytoplasmic, while the R-DNA polymerase was more evenly distributed between nucleus and cytoplasm. Both embryo and trophoblast had the same distributions. The specific activities of both D-DNA and R-DNA polymerases were higher in the embryo than in the trophoblast or decidua (experiment 2). To determine whether levels of DNA polymerase in trophoblast cells doubled TABLE 1 with every doubling of DNA during enTEMPLATE SPECIFICITY OF MOUSE R-DNA POLYMERASE doreduplication, the ratio of DNA polymerase activity per milligram DNA was Expt. Template Activity” 9% No. Activity calculated. This ratio was 7-9 times higher for the D-DNA polymerase and 4-6 times 1 Oligo(dT) .poly(rA) 770 100 higher for the R-DNA polymerase in the Poly(dT) .poly(rA) 1677 218 embryo than the trophoblast (Table 2). Oligo(rU) .poly(rA) 436 57 Poly( rU) poly( rA) Substantial differences were also found to 51 7 Poly(rA) 63 8 exist between the embryo and decidua DNA polymerase : DNA ratios. The specific 2 Oligo(dT) .poly(rA) 5658 100 activity differences between embryo, troPoly(dT) .poly(rA) 8535 151 phoblast, and decidua were larger when Oligo(dT) 39
Incubation
Time (Mid
FIG. 6. Kinetics of the D-DNA and R-DNA polymerase reactions in embryo and trophoblast. The preparations used had been chromatographed on DEAE-cellulose columns. Assay conditions are described in Materials and Methods. (a) D-DNA polymerase; (b) R-DNA polymerase. O-0, Embryo 04, trophoblast.
SHERMAN AND KANC
Mouse Embryonic TABLE
LEVELS OF D-DNA
AND R-DNA
Nuclear
Embryo Trophoblast Experiment 2 Embryo ~e;plmi;~last
Embryo Fe;I$$last
2
POLYMERASEACTIVITY IN ELEVENTH DAY EMBRYO, TROPHOBLAST,AND DECIDUA % Activity
Experiment 1 Embryo Trophoblast
207
DNA Polymerases
Specific activity” (pmoles/min/mg)
Cytoplasmic
Protein
DNA
Embryokrophoblast (or embryo/decidua) Protein*
DNA’
22 23
D-DNA 78 77
Polymerase -d -
-
2.9
7:
:z
R-DNA 56 54
Polymerase 2.4 1.4
71 16
1.7
4.5
D-DNA
Polymerase 170 52 19
4868 541 266
31 8.9
9.0 18.3
Polymerase 5.1 2.6 1.8
145 27 25
2; 2.8
-
R-DNA -
Et:
a Specific activities were calculated on the basis of the protein (Lowry et al., 1951) or DNA (Burton, 1956) content of the initial untreated homogenate. bThe ratio of specific enzyme activity on a per milligram protein basis in the embryo to that in trophoblast or decidua. c The ratio of specific enzyme activity on a per milligram DNA basis in the embryo to that in trophoblast or decidua. d Because high specific activity [3H]dTTP was used in this experiment, substrate concentrations were too low to provide a reliable estimate of the specific enzyme activity.
DISCUSSION
In the course of our experiments, we did not find any qualitative differences in the D-DNA and R-DNA polymerases in mouse embryo, trophoblast, or decidua on the 11th day of gestation. Such differences might be present at other gestation ages. It would, however, be difficult to so expand our studies because of the problems involved in separating trophoblast from decidua at later stages (Sherman, 1972) and obtaining adequate amounts of tissue at earlier stages. Although the nuclear and cytoplasmic D-DNA polymerases were separable under appropriate conditions by glycerol gradient centrifugation, they appeared to cochromatograph on DEAE-cellulose. In contrast, Weissbach et al. (1971) found two peaks of D-DNA polymerase activity when HeLa cell nuclei were chromatographed on
a DEAE-cellulose column: 1540% of the activity passed unadsorbed through the column in the 0.02 M phosphate buffer wash, and the remainder of the activity eluted at higher salt concentrations. Recently, using relatively large quantities of tissue, we observed in nuclear, but not in cytoplasmic, extracts, some D-DNA polymerase activity eluting in the wash fraction from a DEAE-cellulose column (following passage through an initial preparative column). However, even under these conditions, a substantial fraction of the activity was retained on the column. The results presented here suggest that in our hands the nuclear D-DNA and the R-DNA polymerase show a strong affinity for residual endogenous DNA; unless the DNA is removed, this interaction affects the chromatographic and sedimentation profiles of the enzymes. A further complication might
208
DEVELOPMENTAL BIOLOGY
be the possible interconversion of the high and low molecular weight D-DNA polymerases, suggested by the recent studies of Chang and Bollum (1972b) and Hecht (1972). It should, nevertheless, be noted that in any given experiment, regardless of the purification procedure, the sedimentation or chromatographic profiles were the same in the three tissues under study. Weissbach et al. (1971) reported that HeLa nuclear extracts contained both low and high molecular weight D-DNA polymerases while the cytoplasmic extract contained only the latter. Chang and Bollum (1972a), on the other hand, observed that rat liver nuclei contained only the low molecular weight enzyme, while the cytoplasmic extract contained both enzymes. Finally, Hecht (1972) found that in the mouse testis, nuclear extracts contained only the low molecular weight D-DNA polymerase, while the cytoplasmic extract contained only the high molecular weight species. Our results are in close agreement with those of Hecht (1972). The discrepancies may be due to the different species studied, or the method of preparing the nuclear and cytoplasmic extracts. Fry and Weissbach (1973) observed that the R-DNA polymerase of mouse L-cells in culture was largely cytoplasmic. As Table 2 indicates, we have found an equal distribution of the enzyme between nucleus and cytoplasm. The significance of this difference awaits further study. Autoradiographic studies indicate that the S phase in giant trophoblast cells is relatively short and that all giant cells are capable of incorporating 3H-thymidine into acid-insoluble material (Cameron, 1964; Zavarzin et al., 1966; Sherman, unpublished observations). The data in Table 2, however, show that the specific activities of R-DNA polymerase and total D-DNA polymerase in embryo were twice and three times as high, respectively, as in trophoblast. These differences were amplified when calculated on a per milligram of DNA basis (Table 2). The differences in specific
VOLUME34, 1973
activity were not due to cochromatography of inhibitors in the trophoblast preparation or activators in the embryo preparation, since activity was additive when the trophoblast and embryo enzymes were mixed and assayed (unpublished results). If the levels of the DNA polymerases were rising proportionately with the accumulation of DNA in trophoblast giant cells, the ratio of enzyme activity to DNA content should be similar to that observed in embryo cells. On the other hand, if there were no increase in the levels of DNA polymerase as polyploidization occurred, this ratio in the embryo preparation would be expected to be between 13 and 39 times that in the trophoblast preparation, calculated on the basis of 10-30s of the trophoblast cells having undergone polyploidization and reached ploidy levels averaging 256n by the 11th day of gestation (Barlow and Sherman, 1972). The intermediate values obtained for both D-DNA (7-9 times) and R-DNA (4-6 times) polymerases suggest that some increase in the enzyme levels had occurred during polyploidization, but not to the extent required to maintain the enzyme: DNA ratios. If the DNA polymerases studied here are largely responsible for replication of the genome, then relatively low levels of enzyme would appear to be adequate to replicate the greatly enlarged amount of DNA in trophoblast giant nuclei in a reasonably short time. It would then follow that the levels of DNA polymerase present in diploid mouse embryonic cells are far in excess of that required for replication. It must be pointed out that the relationship between the enzymes studied in this report, and in others, to replication in uiuo has yet to be established. Grippo and Lo Scavo (1972) have provided evidence that a new DNA polymerase activity appears in mature Xenopus laevis oocytes in preparation for the onset of replication. Chang and Bollum (1972a) have observed that only the 6-8 S cytoplasmic DNA polymerase activity rises significantly during rat liver
SHERMAN AND KANG
Mouse Embryonic
regeneration, leading them to propose that this enzyme is involved in DNA replication in c&o. Fansler and Loeb (1969) observed a redistribution of DNA polymerase from cytoplasm to nucleus during embryogenesis in the sea urchin. The cell types in the present study are either (a) replicating and dividing, (b) replicating but not dividing, or (c) neither replicating nor dividing. Yet we have not observed in any of these cases either the presence of a unique enzyme, any striking difference in the relative proportions of the enzymes, or any significant variation in the distribution of the enzymes between nucleus and cytoplasm. It would appear from our experiments that the nuclear and cytoplasmic D-DNA polymerases are coordinately controlled, both being present at high levels in actively dividing cells, and at lower levels in nondividing cells. The role of the R-DNA polymerase in in uiuo DNA synthesis is at present ambiguous. Fry and Weissbach (1973) have carried out a detailed study on an enzyme in mouse L cells with properties very similar to those described here. Although this enzyme is not a terminal transferase and can utilize a number of artificial templates (see also Table l), in vitro conditions have not yet been found which will enable the enzyme to use bacteriophage, viral, or natural mRNA as template (Fry and Weissbach, 1973). Nevertheless, none of our data refutes Temin’s hypothesis that an R-DNA polymerase, perhaps the enzyme described here, plays a role in cell differentiation (Temin, 1971). If this were the case, however, the presence of significant amounts of the same enzyme in decidua, a maternal, nondividing tissue would have to be explained. We wish to thank Ms. N. Chew, Dr. A. Weissbach, and especially Dr. M. Fry for their help. REFERENCES ALDEN, R. H. (1948). Implantation of the rat egg. III. Origin and development of primary trophoblast cells. Amer. J. Anat. 83. 143-182.
DNA Polymeruses
209
ATLAS,M., BOND, V. P., and CRONKITE, E. P. (1960). DNA synthesis in the developing mouse embryo studied with tritiated thymidine. J. Histochem. Cytochem. 8, 171-181. BARLOW, P. W., and SHERMAN, M. I. (1972). The biochemistry of differentiation of mouse trophoblast: studies on polyploidy. J. Embryol. Exp. Morphol. 27, 447-465. BOLDEN, A., FRY, M., MULLER, R., and WEISSBACH,A. (1972). The presence of a polyriboadenylic aciddependent DNA polymerase in eukaryotic cells. AFC~. Biochem. Biophys. 153, 26-33. BURTON, K. (1956). A study of the conditions and mechanism of the diphenylamine reaction for the calorimetric estimation of the deoxyribonucleic acid. Biochem. J. 62, 315-323. CAMERON, I. L. (1964). Is the duration of DNA synthesis in somatic cells of mammals and birds constant? J. Cell Biol. 20, 185-188. CHANG, L. M. S., and BOLLUM, F. J. (1971). Low molecular weight deoxyribonucleic acid polymerase in mammalian cells. J. Biol. Chem. 246,5835-5837. CHANG, L. M. S., and BOLLUM, F. J. (1972a). Variation of deoxyribonucleic acid polymerase activities during rat liver regeneration. J. Biol. Chem. 247, 7948-7950. CHANG, L. M. S., and BOLLUM, F. J. (197213). Antigenie relationships in mammalian DNA polymerase. Scence 175, 1116-1117. CHAPMAN, V. M., ANSELL, J. D., and MCLAREN, A. (1972). Trophoblast giant cell differentiation in the mouse: expression of glucose phosphate isomerase (GPI-1) electrophoretic variants in transferred and chimeric embryos. Deuelop. Biol. 29, 48-54. FANSLER, B., and LOEB, L. A. (1969). Sea urchin nuclear DNA polymerase. II. Changing localization during early development. Exp. Cell Res. 57, 305-310. FRIDLENDER, B., FRY, M., BOLDEN, A., and WEISSBACH, A. (1972). A new synthetic RNA-dependent DNA polymerase from human tissue culture cells. Proc. Nat. Acad. Sci. U.S. 69, 452-455. FRY, M., and WEISSBACH,A. (1973). The utilization of synthetic DNA templates by a new DNA polymerase from cultured murine cells. J. Biol. Chem. 248, 2678-2683. GEARHART, J. D., and MINTZ, B. (1972). Glucosephosphate isomerase subunit-reassociation tests for maternal-fetal and fetal-fetal cell fusion in the mouse placenta. Develop. Biol. 29, 55-64. GRAHAM, C. F. (1973). The cell cycle during mammalian development. In “The Cell Cycle in Development and Differentiation” (M. Balls and F. S. Billett, eds.), pp. 293-310. Cambridge Univ. Press, London and New York. In press. GRIPPO, P., and Lo SCAVO,A. (1972). DNA polymerase activity during maturation in Xenopus laevis oocytes. Biochem. Biophys. Res. Commun. 48, 280285.
210
DEVELOPMENTAL BIOLOGY
HECHT, N. B. (1972). The interconvertibility of mouse DNA polymerase activities derived from the nucleus and cytosol. J. Cell Biol. 55, 109a. HUNT, C. V., and AVERY, G. B. (1971). Increased levels of deoxyribonucleic acid during trophoblast giant cell formation in mice. J. &prod. pert. 25, 85-94. LOWRY, 0. H., ROSEBROUGH,N. J., FARR, A. L., and RANDALL, R. J. (1951). Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265-275. MAIA, J. C. C., ROUGEON, F., and CHAPEVILLE, F. (1971). Chick embryo poly(rA:dT)-dependent DNA polymerase. FEBS Letr. 18, 130-134. SCHLABACH, A., FRIDLENDE~ B., BOLDEN, A., and WEISSBACH, A. (1971). DNA-dependent DNA polymerases from HeLa cell nuclei. II. Template and substrate utilization. Biochem. Biophys. Res. Com-
mun. 44,879-885. SHERMAN, M. I. (1972). The biochemistry of differentiation of mouse tropboblast: alkaline phosphatase.
Develop. Biol. 27, 337-350. SHERMAN, M. I., WALKER, P. M. B., and MCLAREN, A. (1972). Mechanism of accumulation of DNA in Nature (London) giant cells of mouse trophoblast. New Biol. 238, 175-176. SLATER, I., and SLATER, D. W. (1972). DNA polymerase potentials of sea urchin embryos. Nature (London) New Biol. 27, 81-84. STAVRIANOPOULOS,J. G., KARKAS, J. D., and CHARGAFF,
VOLUME 34, 1973
E. (1971). Nucleic acid polymerases of the developing chicken embryo: a DNA polymerase preferring a hybrid template. Proc. Nat. Acad. Sci. U.S. 68, 2207-2211. TEMIN, H. M. (1971). The protovirus hypothesis. J. Nut. Can. Inst. 46: III-VII. WARD, D. C., HUMPHREYS, K. C., and WEINSTEIN, I. B. (1972) Synthetic RNA-dependent DNA polymerase activity in normal rat liver and hepatomas.
Nature (London) 237, 499-503. WEISSBACH, A., SCHLABACH, A., FRIDLENDER, B., and BOLDEN, A. (1971). DNA polymerases from human cells. Nature (London) New Biol. 231, 167-170. ZAVARZIN, A. A., SAMOSHKINA, N. A., and DONDUA, A. K. (1966) Synthesis of DNA and kinetics of cells in early embryogenesis of mice. Zh. Obshch. Biol. 27,
697-709. ZYBINA, E. V. (1960). Sex chromatin in the trophoblast of early white rat embryos. Dokl. Akad. Nauk.
SSSR 130, 633-635. ZYBINA, E. V. (1961). Endomitosis and polyteny of trophoblast giant cells. Dokl. Akad. Nauk. SSSR 140, 1177-1180. ZYBINA, E. V. (1963). Cytophotometric determination of polyploidization of the cells of the trophoblast. Dokl. Akad. Nauk. SSSR 153, 1428-1431. ZYBINA, E. V., and MOS’YAN, I. A. (1967). Sex chromatin bodies during endomitotic polyploidication of trophoblast cells. Tsitologiya 9, 265-272.