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Sea urchin nuclear DNA polymerase
Copyright 0 I972 by Academic Press, Inc. All rights of reproduction in any form reserved
Experimental Cell Research 75 (1972) 433-441
SEA URCHIN
NUCLEAR
IV. Reversible Association
DNA POLYMERASE of DNA Polymerase with
Nuclei During the Cell Cycle’ B. FANSLER and L. A. LOEB Institute for Cancer Research, Fox Chase, Philadet’phia, Pa 19111, USA
SUMMARY We have taken advantage of the natural synchrony of sea urchin embryos during early development to study the relationship between DNA polymerase activity and nuclear DNA replication. Nuclei were isolated from early embryos (2-16 cells) in various phases of the division cell cycles. The polymerase activity found in nuclei isolated from S phase cells was 5 to lo-fold greater than that in similar fractions isolated from mitotic or G 2 embryos. As DNA synthesis is completed, the polymerase activity drops to a level slightly higher than the comparable stage before S. The ratio of DNA polymerase activity to in vivo synthesized DNA is consistently highest at the beginning of S. If mitosis is prolonged, by the isotope effect of D,O on mitotic spindle proteins, there is a corresponding delay in the increase of nuclear polymerase activity and DNA replication. We interpret these results to mean that DNA polymerase attaches to the chromosomes as they complete mitosis, functions in DNA synthesis, dissociates from them and leaves the nucleus when DNA replication is completed. This translocation is repeated with each cell cycle.
Early development of sea urchin embryos is characterized by exponential cell division with accompanying DNA replication. Our previous studies [l, 21 have shown a high level of DNA polymerase activity in unfertilized eggs and this amount of activity per embryo does not change during early development. In further studies we made a direct comparison between the DNA polymerase activity of isolated nuclei and the sea urchin embryos from which they were derived. Using embryos taken at various times between fertilization and blastula (18 h at 15”(Z), we isolated nuclei and a cytoplasmic fraction and prepared a whole embryo homogenate. DNA polymerase activity was assayed on each of these preparations. As development proceeds 1 Paper III in this series is Ref. 3. 29 -
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the activity in the nuclear fraction increases, that of the cytoplasmic decreases and the whole embryo activity remains constant. By the blastula stage, when cleavage has slowed considerably, the majority of the DNA polymerase activity is associated with the nuclei. Studies in which DNA polymerase was purified to homogeneity from embryos briefly exposed to radioactive amino acids [3] indicate that the synthesis or turnover of this enzyme during early development is no greater than most other proteins. These studies were supported by the work of Fry & Gross [4] who showed that only about 20% of the sea urchin embryo’s total protein undergoes turnover in 24 h after fertilization. Since little polymerase is synthesized during early development, we can presume that the progressive increase in nuclear polymerase activExptl Cell Res 7.5(1972)
434
B. Fader
& L. A. Loeb
itv renresents a molecular transfer of the enzyme from the cytoplasm to the nucleus. However, we do not know when during the cell cycle the enzyme becomes associated with the nucleus or what happens to it after that. We now report on experiments in which nuclei were isolated from synchronously dividing embryos at several points during the cell cycle. The nuclear fractions and chromatin isolated from them were assayed for DNA polymerase activity. The results suggest that DNA polymerase attaches to the chromosomes as mitosis is completed, and then dissociates from the chromosomes. -d
-A-
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MATERIALS The sea urchins, Strongylocentrotus purpuratus were purchased from Pacific Bio-Marine, Venice, Calif. Homogeneous sea urchin DNA polymerase was prepared from nuclei of S. purpuratus blastulae (fraction VIII) [S]. Unlabeled deoxynucleoside triphosphates were purchased from CalBiochem. c@P-thymidine triphosphate was purchased from International Chemical and Nuclear Coruoration. Citv of Industrv. Calif. Calf thymus DGA was &qui;ed from Woi: thington Biochemical Corp. Deuterium water (D,O) waspurchased from Sigma Chemical Co. ‘Max&n& activated’ calf thymus DNA was prepared as described by Loeb [5]. Artificial sea water was made from distilled water and salts to the following concentration: 0.446 M NaCl, 0.0255 M MgCl,, 0.5182 M MgSO,, 0.009 M KCI. 0.0022 M NaHCO,, and 0.097 M CaCl,. This was us&d for all experim&ts unless otherwise indicated.
METHODS Preparation of hatching enzyme Sea urchin embryos were grown to the hatching stages, about 18 h at 15°C. When 95 % had visibly lost their fertilization membranes the culture was cooled to 4°C and the embryos filtered from the sea water containing the hatching enzyme. This can be kept chilled uv to 6 h without loss of activitv. Alternativelv. after filtration (Millipore, 0.45 pm), _the solution can be stored frozen at -20°C for several months. It is necessary to concentrate the thawed solution about 5 times by Diaflo ultrafiltration, UM-1 filter, as enzyme activity is lost upon freezing.
Protocol for synchrony experiments Sea urchin eggs were shed by injecting0.5 M KCl. The eggs were collected into sea water, the sperm were Exptl Cell Res 75 (1972)
collected ‘dry’ into weighing boats. Washing and fertilizing the eggs were as described by Mazia et al. 161.Eggs from one urchin (610 ml packed; about 1 in 50 females yield enough eggs for these experiments) were fertilized in 100-200 ml sea water at 15°C. Only batches in which 98 % of the eggsfertilizedwithin2min were used in these experime&& After 5 min, 6-8 vol of sea water containing hatching enzyme were added. Membranes were not visible 1 to 2 h after adding the enzyme. At a time between 50 and 130 min after fertilization, SH-thymidine (3H-TdR, spec. act. 6.7 Ci/mM) was added to the culture. The embryos were then allowed to develop to the desired stage. The precision of the synchrony was monitored at frequent intervals by microscopic examination. The timing of DNA synthesis during the cell cycle was monitored by measuring the incorporation of 3H-TdR into whole embryos. 1.0 ml aliquots, in duplicate, are removed from the developing culture (about lo* embryos) and the amount of 3H-TdR incorporated was determined by the method of Hinegardner et al. [7].
Nuclear isolation Aliquots for nuclear isolation (l-2 x 106) embryos were immediately mixed with 2 vol of a sea water ice slush to stop development. All procedures were performed at 04°C and all centrifugations are 2 500 g for 10 min unless otherwise stated. The embryos were centrifuged from the iced sea water, put into 30 ml Corex round bottom tubes and washed twice in 25 ml of 1.0 M dextrose and once in 25 ml of a hypotonic solution of 0.15 M sodium chloride with 0.015 M sodium citrate. The pellet was suspended in 12 ml of the hypotonic solution and was then dispersed by forcing the suspension 3 times through -a syringe number 20 needle. An equal volume of 2.0 M sucrose was thoroughly mixed- with the suspension after which it was centrifuged at 14 500 g for 30 min. Care was taken to decant the supernatant without contaminating the pellet with the floating yolk material. After draining the tubes and removing excess sucrose solution, the nuclear pellets were dispersed in 0.5 ml of solution E: 20 % glycerol, 0.02 M potassium phosphate buffer, pH 7.8, 4.0 mM reduced glutathione and 0.4 mM potassium versenate and sonicated intermittently at O-1°C. This is referred to as the nuclear fraction. When this method was used on cells in mitosis with no apparent nuclear membrane, the recovery of nuclear material as judged by microscopic examination and recovery of radioactive DNA in the nuclear fraction was not significantly different from nuclei in other phases of the cell cycle.
Chromatin preparation Chromatin isolation from nuclei was carried out in 20 % glycerol since polymerase requires polyglycols for stability [5]. The nuclei were suspended in 20 % glycerol, broken by intermittent sonication at 0-1°C and dialysed 18 h against 2 1of 20 % glycerol. Particulate material was removed by centrifugation at 2 500 g. The supernatant was then centrifuged at 100 000 g in an SW 40.1 rotor for 90 min. The resulting
Sea urchin nuclear DNA polymerase. IV pellets were suspended by homogenization in 1.0 ml of solution E. The nm ratio 280/260 is about 0.8.
DNA polymerase activity The assay measures the incorporation of [a+P]thymidine triohosohate into an acid insoluble nroduct. The reaction mixture, in a total volume of-O.3 ml, contains 20 % glycerol, 25 pmoles Tris-maleate buffer, pH 8.0; 4.5 pmoles MgC12; 0.3 pmole 2-mercaptoethanol; 10 nmoles each of dATP, dCTP, dGTP and [@2p]dTTP (about 3 x 10” dpm/nmole); 177 nmoles of ‘activated’ calf thymus DNA (expressed as mononucleotide and O.Oj ml of nuciea; fraction). After incubation 0.1 ml of 1.0 mg/ml denatured DNA was added and the reaction was stopped with 2.0 ml of cold 0.5 N perchloric acid containing 0.01 M sodium pyrophosphate. After standing for 10 min at O”C, the precipitate was pelleted by centrifugation at 5 000 g for 20 min. The pellets were dissolved in 0.5 ml of 0.2 M sodium hydroxide and precipitated with 1.Oml of the cold perchlorate-sodium pyrophosphate solution. After 10 min at O”C, 3.0 ml of cold water were added and the precipitate collected on a glass fiber filter (Whatman GF/C, 2.4 cm). The filter was washed once with 0.5 ml of the uerchlorate-sodium pyrophosphate solution, twice with 10 ml cold water, once with 2.0 ml of 95 % ethanol, transferred to a glass vial, dried and the radioactivity measured [2]. Protein was determined by the method of Lowry et al. [8] using crystalline pancreatic ribonuclease as g standard. Protein fractions containing interfering substances such as glutathione, 2-mercaptoethanol, MgCI, and Tris-maleate were first isolated by precipitation with cold perchloric acid.
The protocol for these experiments was to fertilize the eggs from one urchin and allow them to develop synchronously in the presence of 3H-TdR. The timing of DNA synthesis during the cell cycle was monitored by frequently taking small aliquots of embryos to determine 3H-TdR incorporation into whole embryos. At intervals of 5 or 10 min during one cell cycle, larger aliquots of the culture were taken for nuclear isolation. These nuclei, or crude chromatin isolated from them, were assayed for DNA polymerase activity using a-32P-TdRtriphosphate as the labelled substrate. The 3H-DNA which is in the nuclear fractions used as the source of polymerase also precipitates in the assay procedure. So the ratio of polymerase activ-
1.4
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P I' ' '!
-. 1.2
-- 0.12 4--- 0.08 --0.04 e--- 0
0.6
-04
90
RESULTS
435
120
150
180
210
240
270
Fig. I. Abscissa: time after fertilization (min); ordinate: (left) O-O, aH-TdR incorporation; (right) O---O, DNA polymerase activity (cpm 52P)/thymidine incor-
poration@pm$H).
Ratio of DNA polymerase activity (““P cpm) to thymidine incorporation (3H cpm) in nuclei isolated from embryos during three cell cycles. In expts 1, 2 and 3, 10, 6 and 10 ml of eggs respectively were fertilized and the fertilization membranes removed with hatching enzyme as described in Methods. In expt 1, 0.1 mCi of 3H-TdR was added 50 min after fertilization. In expt 2, 0.3 mCi SH-TdR was added 130 min after fertilization. In expt 3, 0.2 mCi SH-TdR was added 110 min after fertilization. Nuclear isolation, polymerase assays and 3H-TdR incorporation into whole embryos were determined as in Methods. In all experiments, polymerase assays were incubated 30 min at 37°C.
ity, i.e. 32P,to in vivo synthesized DNA, i.e. 3H, can be obtained directly on the same sample. This ratio is very reliable since it is not dependent upon variations in recovery of nuclear material at different points across the Exptl Cell Res 75 (1972)
436
B. Fader
& L. A. Loeb
OL Fig. 2. Abscissa: time after fertilization (min); dinate:
(left)
O-O, 3H-TdR incorporation (cpm x 1O-5); O---O nmoles TM a2P/mg protein. DNA polymerase activity per mg protein in nuclei isolated from embryos during the 4-8 cell cycle. Details are given in the caption to fig. 1 as expt 2. Synchrony refers to the incorporation of aH-TdR into DNA by the whole embryo.
(right)
cell cycle. A sharp increase in the ratio is significant as an indication that DNA polymerase has become associated with the nuclei. Fig. 1 shows the ratio of DNA polymerase activity to thymidine incorporation from three experiments over three cell cycles from 2 to 16 cells. The synchrony curves (thymidine incorporation) show that the cells were well synchronized in each experiment; having a cell cycle of 60 min with DNA synthesis occupying at most 20 min. (There is no G 1 in these cells. DNA synthesis begins in midtelophase and ends in early G2). During mitosis (140, 200, 260 min) the ratio is very low. However, with the beginning of DNA synthesis in each cycle (150, 210, 270 min) the ratio increases several-fold to its highest value. In experiment 3, nuclei were isolated over two S periods to show that this pattern of change in the ratio is consistent from one cell cycle to the next. These results indicate that the DNA polymerase becomes associated with the nucleus at the very beginning of the DNA synthetic period. As DNA synExptl Cell Res 75 (1972)
thesis is completed, the ratio decreases (160, 170, 220, 230 and 280 min), but it is difficult to determine from these data whether it was due to the increase in thymidine incorporation (tritium counts) alone, or this increase in tritium counts coupled with an actual decrease in polymerase activity. In other words, does the DNA polymerase remain attached to the chromosomes through mitosis or does it dissociate from them at the end of S? Fig. 2 gives the DNA polymerase activity of nuclei isolated from embryos at various phases of a 4-8 cell cycle. This is the specific activity data (polymerase activity per mg protein) of expt 2 in fig. 1. In contrast to the total embryo DNA polymerase activity which varies only about one-fold during the cell cycle [9], activity in the isolated nuclei increases ten-fold as the cells begin DNA synthesis. In this, and the other 4-8 cell experiment, the highest polymerase activity is in nuclei from mid-S, but in most other experiments it has been at the beginning of S. In all the experiments, however, the nuclear DNA polymerase activity decreases sharply as the cells complete DNA synthesis, reaching a minimum very soon thereafter. The most direct interpretation of these results is that DNA polymerase leaves the nucleus as replication is completed. It is to be noted that in all the experiments the polymerase activity after DNA synthesis is slightly higher than at the comparable stage before. It is possible that the increase in specific activity of polymerase in nuclei at the start of replication is simply unbound enzyme selectively trapped in the nucleus by the reformation of the nuclear membrane which occurs in early telophase. This is improbable because the nuclear membrane reforms out from the chromosomes in a manner which seemingly excludes most of the cell cytoplasm [lo]. Nevertheless, to show that polymerase is associated with chromatin, a crude fraction
Sea urchin nuclear DNA polymerase. IV was preared from the nuclei isolated over a 4-8 cell cycle. To insure the stability of the polymerase, the nuclei were suspended in 20 % glycerol and broken by sonication. After dialyzing and centrifuging out particulate matter, the chromatin was pelleted from the 20% glycerol. If this pellet is resuspended by homogenization in 20% glycerol and repelleted, there is only 5 % loss of polymerase activity. We assumed from this that the enzyme which is associated with the chromatin pellet must actually be bound to the DNA. The results of polymerase assays in these chromatin fractions are presented in fig. 3. The polymerase activity per mg protein and the ratio of polymerase activity to thymidine incorporation is low during mitosis and reach a maximum at the very beginning of S, 210 min, as in the isolated nuclei. When the DNA polymerase assays were done on these chromatin fractions without any added primer DNA, the pattern of polymerase activity over the cell cycle was the same with only about 10 Y0 of the enzyme activity. The polymerase activity also is highest at the start of DNA replication. But apart from this, the results are somewhat difficult to interpret because the amount of enzyme activity declines precipitously during DNA synthesis and actually reaches a minimum slightly before it is completed. This may be explained to some degree by the sonication used to break the nuclei; for example, the complex of chromatin and DNA polymerase may be particularly delicate and susceptible to shock during S phase. The conclusions from these chromatin assays must be drawn with some reservations. Nevertheless, because the pattern of polymerase activity over the cell cycle closely resembles that of the nuclear fractions, and because the highest ratio of polymerase activity to thymidine incorporation is at 210 min, the very beginning of S, it seems reasonable to assume that at least part of the DNA
437
Fig. 3. Abscissa: time after fertilization (min); ordinate: (left) O-O, SH-TdR incorporation (cpm x 10m4); A--A (right) (outer) DNA polymerase activity (cpm TM 32Pincorporation)/thymidine incorporation (cpm 3H); (inner) nmoles TM 82P/mg protein. DNA polymerase activity in chromatin prepared from nuclei isolated at different times during the 4-8 cell cvcle. Nine ml of eaas were fertilized and the fertilization membranes k-moved with hatching enzvme as described in Methods. 0.2 mCi 3H-TdR was added at 100 min. Nuclear isolation, chromatin isolation, polymerase assays and 3H-TdR incorporation into whole embryos were determined as in Methods. Polymerase assays were incubated for 1 h at 37°C.
polymerase entering the nucleus at the beginning of DNA synthesis becomes bound to the chromosomes. Is this decrease in polymerase activity at the end of S actually due to a loss of enzyme, or possibly some inhibitor which appears at that time? In order to check for enzyme inhibition, purified sea urchin DNA polymerase was mixed with nuclei having the lowest activity and we looked for additivity of the enzyme activities. For example, in an experiment in which the activity for the nuclear fraction (time 250, expt 3, fig. 1) was 250 pmoles TMP incorporated, and the purified enzyme incorporated 120 pmoles, together the activity was 390 pmoles. This indicates additivity between the purified and nuclear enzyme activities thus ruling out any readily soluble inhibitor present in excess under the conditions of the in vitro assay. Exptl Cell Res 75 (1972)
438 B. Fader & L. A. Loeb
Fig. 4. Abscissa: time after fertilization (min); ordinate: (left) 3H-TdR incorporation (cpm x 10--4). O-O, D,O treated embryos; O-O, untreated embryos; (right) o---n, nmoles TM 52P/mg protein. Effect of D,6 on DNA synthesis and nuclear DNA polymerase activity. Seven ml of eggs were fertilized and thefertilization membranes removed with hatching enzyme as in Methods. After 90 min the embryos were allowed to settle and the sea water volume reduced to 125 ml. 0.1 mCi of SH-TdR was added at 130 min. A 10 ml portion of the culture was removed and allowed to develop normally. From this, 0.5 ml synchrony aliquots were taken at times indicated in order to monitor normal DNA synthesis. At 180 min after fertilization 15 ml of the culture were taken for nuclear isolation. Immediately, 100 ml of D,O sea water were added to the remaining 100 ml of the culture to give a 50 % D,O final concentration. One ml synchrony aliquots were taken from time 180 to time 200. At 190 and 210 min, 30 ml aliquots were taken for nuclear isolation. Immediately after the 210 min aliquot was taken, 1 440 ml sea water were added to give a final D,O concentration of 5 % which allows the embryos to develop normally. After this time, 10 ml synchrony and 300 ml nuclear isolation aliquots were taken. All procedures for nuclear isolation and polymerase assays are as in Methods. Assays were incubated 30 min at 37°C.
We then wanted to know if this pattern of increase and decrease in DNA polymerase activity over the cell cycle is unique to this enzyme, or might it be common to other enzyme activities under these conditions. To answer this, nuclear fractions were assayed for three other enzyme activities, DNase, thymidine kinase, and lactic acid dehydrogenase; chosen because of their widely diverse functions with respect to cellular metabolism. While there are variations in these enzyme Exptl Cell Res 75 (1972)
activities over the cell cycle, none of the three in any way shows the consistent pattern of change which is observed for nuclear DNA polymerase activity. It appears, then, from these studies that the increase and decrease of nuclear DNA polymerase activity with the onset and completion of each S period is a property of this enzyme. To show a correlation between the rise in polymerase activity and the beginning of DNA synthesis, synchronized embryos were reversibly stopped in metaphase. The delay was accomplished by adding an equal volume of 100% deuterium oxide (DzO) sea water to a culture of embryos in prophase. Gross & Spindel [11] have shown that echinoderm embryos in 50% D,O sea water will not develop past metaphase, presumably because of an isotope effect preventing the mitotic spindle subunits from dissociating. This inhibition is reversed within 1 min of lowering the D,O concentration to 5 %. Embryos were in 50 Y0 D,O for a 30 min period from 180 to 210 min post-fertilization and then diluted into 9 vol of sea water without DzO. Aliquots were taken and nuclei isolated from them at the points indicated in fig. 4. In addition, a small amount of the culture not exposed to D,O was allowed to develop in order to monitor normal DNA synthesis. Fig. 4 shows the DNA polymerase activity of nuclei isolated from cells during and after D,O inhibition. DNA synthesis is delayed about 25 min, from time 220 to about 245 min after fertilization. Nuclear DNA polymerase activity decreases through the period of D,O treatment and after, until, at mid-telophase it shows the characteristic marked increase. We conclude that when mitosis is extended, there is a proportional delay in the association of DNA polymerase with the chromosomes and in the subsequent DNA synthesis.
Sea urchin nuclear DNA polymerase. IV DISCUSSION We have presented evidence which suggests that the DNA polymerase in early developing sea urchin embryos becomes associated with the nucleus at the very beginning of DNA synthesis and subsequently leaves the nucleus as replication is completed. Furthermore, at least part of the nuclear polymerase attaches to the chromatin during S, although the association may be rather delicate and easily disrupted. This pattern of reversible association is repeated in each cell cycle with the DNA polymerase presumably residing in the cytoplasm during G2 and mitosis. The decrease in polymerase activity at the end of S is not due to a soluble inhibitor present in excess. If mitosis is prolonged by the isotope effect of D,O on mitotic spindle proteins, there is a corresponding delay in the increase of nuclear DNA polymerase activity and DNA replication. These results should be considered in the context of the nucleo-cytoplasmic events during the cell cycle. At the second, third and fourth cleavages only 5-15 % of the unfractionated embryo’s total DNA polymerase activity can be recovered in isolated nuclei. This enzyme activity and what seems more significant, the activity associated with chromatin shows an abrupt several-fold increase at the beginning of each S. In the sea urchin embryo this is the point in mitosis when the chromosomes begin to extend after being in intimate contact with the cytoplasm. Presumably, as the chromosomes unwind, many points on the DNA become available for enzyme attachment. Loeb [12] has estimated, on the basis of the polymerase activity bound to a crude chromatin preparation, from the amount of activity of the homogeneous enzyme, and the amount of DNA per embryo, that one DNA polymerase molecule could bind for each two to three thousand nucleo-
439
tide pairs. The binding points on the DNA are presumably 3’-OH groups. Loeb [5] has shown that DNA treated with pancreatic DNase to produce a 3’-OH is an excellent primer for sea urchin polymerase while DNA digested with micrococcal nuclease to give 3’-PO, will not prime. These binding points, could result from the activity of an endonuclease similar to that described by Burgoyne et al. [13]. They reported a calciumdependent endonuclease activity which enables DNA polymerase in rat liver nuclei to use the nuclear DNA as primer-template. After DNA synthesis is completed, the DNA polymerase activity of both the nuclear and chromatin fractions decreases. Since the polymerase activity of the whole embryo remains fairly constant throughout the cell cycle, and indeed, through early development up to gastrulation, we have concluded that this decline in activity is enzyme leaving the chromosomes and the nucleus. However, the possibility of some enzyme inhibitor appearing at the end of S cannot be completely excluded. This reversible association of DNA polymerase with the nucleus during the mitotic cycle was unexpected since it creates an apparent paradox with the results reported earlier from this laboratory [2], that polymerase activity accumulates in the nucleus as cleavage progresses. Fig. 2 shows that polymerase activity after DNA synthesis is slightly higher than at the comparable stage before S. We found this in all the experiments and interpret it to be residual DNA polymerase which remains with the chromosomes when they condense at prophase. This residual activity could accumulate with each cell division during early development and is sufficient to account for the translocation of polymerase activity into nuclei which was observed in the previous study [2]. These results correlate well with Mazia’s Exptl Cell Res 75 (1972)
440 B. Fader & L. A. Loeb [14] observations. In different partially synchronized populations, he counted the number of sea urchin embryos which were in mitosis and measured the DNA polymerase activity associated with a crude chromatin fraction prepared from them. The polymerase activity was highest in chromatin from cells which had just finished mitosis. He proposed that the enzyme attached to the chromosomes at the end of mitosis, replicated the DNA and then dissociated from them before the next mitosis. Other studies [15, 161 on cells which were synchronized by chemically blocking the DNA synthesis with 5-fluoro-2’deoxyuridine showed a small redistribution of DNA polymerase activity from cytoplasmic to nuclear fraction upon release of the block. Friedman [ 171, using HeLa cells, synchronized them by prolonged exposure to amethopterin and adenosine to induce a thymidineless condition. He then induced DNA synthesis by the addition of thymidine and found that nuclear DNA polymerase activity increased as the cells went into S and decreased as it was finished. Lindsay et al. [18] could not demonstrate the same effect in L cells, synchronized with aminopterin. Madreiter et al. [19], also using L cells, synchronized them mechanically in order to avoid the physiological imbalance of blocking agents. They showed a rise in the level of DNA polymerase activity in a particulate fraction as DNA synthesis occurred. They also looked at DNase activity associated with the same fraction and found little variation over the cell cycle. Howell & Hecht [20] have reported on the DNA polymerase and polynucleotide ligase activities during the mitotic cell cycle of the naturally synchronous LiEium microspore cells. They looked only at the whole cell supernatant fraction after high speed centrifugation. There was no significant fluctuation in DNA polymerase activity during the cell cycle, however, there was a sharp Exptl Cell Res 75 (1972)
increase in polynucleotide ligase activity before S which then decreased during S and subsequently disappeared. They suggest that in this system, the amount of ligase activity may have some controlling influence in DNA synthesis. Each of these studies has the disadvantage of either having to establish synchrony by chemical blocks, the total effects of which are unknown, or mechanically choosing cells in one phase of the cell cycle and having only partial synchrony. In the case of Lilium microspores, the study unfortunately included only enzyme activity in whole cell supernatants. It therefore becomes difficult to compare them directly with our studies, but generally they suggest that nuclear DNA polymerase activity is high as DNA is replicated. Friedman’s work further suggests that the enzyme leaves the nucleus after S. None of them, however, establishes any real correlations between naturally induced DNA synthesis and the level of nuclear DNA polymerase activity. With respect to the control of DNA synthesis, the early developing embryo represents an entirely different situation from a fully differentiated eucaryotic cell. The embryonic cells are dividing rapidly and need a large amount of enzyme to replicate their DNA. On the other hand, differentiated cells generally function in their individual ways, in most tissues they seldom divide and do not require large amounts of DNA polymerase. There is evidence which indicates that in some systems the presence or absence of polymerase activity may be one of the primary factors controlling replication. Stockdale 1211 has shown a very close correlation between the decline of DNA polymerase activity and the lack of ability of multinucleated muscle cells to synthesize DNA or divide. He demonstrated a one hundred-fold decrease in the amount of enzyme activity as the mononucleated cells fused to form the multi-
Sea urchin nuclear DNA polymerase. IV nucleated non-dividing skeletal muscle cells. He was not able to determine if the sharp decrease was due to a loss of the enzyme or an accelerated inactivation. Conversely, when lymphocytes are stimulated with phytohemagglutinin there is a long delay before the cells divide. Loeb et al. [22] have shown that during this delay the cells must synthesize DNA polymerase, after which there is then an excellent correlation between the amount of polymerase activity and the rate of DNA synthesis [23]. There are systems, however, where DNA synthesis does not seem to be dependent upon the level of DNA polymerase; for example, in regenerating rat liver DNA synthesis ceases, but the level of DNA polymerase activity remains high [24], indicating that the cessation of DNA synthesis in this system does not depend upon the loss of enzyme activity. The control of DNA synthesis in the sea urchin embryo does not depend upon the synthesis of new DNA polymerase. There is a high level of activity in the unfertilized egg which persists through the blastula stage. Instead, the control mechanism may involve some step in the transfer process of the enzyme from the cytoplasm to the nucleus; possibly the availability of enzyme attachment sites on the chromosome. We can now imagine a series of events as follows. A molecule of DNA polymerase, which has been synthesized during oogenesis and stored in the cytoplasm, comes in contact with a chromosome unwinding from its mitotic configuration. The enzyme attaches, presumably to a 3’-OH DNA terminus which could result from the action of an appropriate endonuclease. The polymerase participates in DNA replication, dissociates from the chromosome and probably leaves the nucleus altogether. A residual polymerase activity remains associated with the chromosomes through mitosis and accmulates in the nucleus with each cell division. This translocation is
441
repeated with each cell cycle, as enough enzyme is made during oogenesis in the sea urchin to accommodate the rapid DNA synthesis through the blastula stage. We thank Dr Jack Schultz for generous counsel. and dedicate this paper to his memory. This investigation was suooorted by USPHS arants CA-l 1524 and CA-l 1835 from NIH and gram GB18419 from NSF. Support was also derived from grants to this Institute: LJSPHS, CA-06927 and RR05539, and an appropriation from the Commonwealth of Pennsylvania.
REFERENCES 1. Loeb, L A, Fansler B, Williams R & Mazia, D, Exptl cell res 57 (1969) 298. 2. Fansler, B & Loeb, LA, Exptl cell res 57 (1969) 305. 3. Loeb, L A & Fansler. B. Biochim bioohvs . < acta ’ ’ 217 (1970) 50. 4. Fry, B J & Gross P R. Dev biol21 (1970) 125. 5. Loeb L A, J biol them 244 (1969) l672.’ 6. Mazia, D, Mitchison, J M, Medina, H & Harris, P J, J biophys biochem cytol 10 (1961) 467. 7. Hinegardner. R T. Rao. B & Feldman. D E. Expti cell res 36 (1964) 53. 8. Lowry, 0 H, Rosebrough, N J, Farr, A L & Randall, R J, J biol them 193 (1951) 265. 9. Loeb, L A, Fansler, B & Salter J P, Symposium on uptake of informative molecules by living cells (ed L Ledoux). North-Holland, Amsterdam and London (1970). 10. Mazia D, The cel1 (ed J Brachet & A E Mirsky) vol. 3, ^_ DU. 77-412. Academic Press. New York and London (1961). Il. Gross, P R & Spindel, W, Ann NY acad sci 90 (1960) 500. 12. Loeb, L A, Nature 226 (1970) 448. 13. Burgoyne, .L A, Waqar, M A‘& Atkinson, M R, Biochem biophys res comm 39 (1970) 254. 14. Mazia, D, Cell nucleus, pp. 15-25. Taylor & Francis, London (1966). 15. Gold, M & Helleiner, C W, Biochim biophys acta 80 (1964) 193. 16. Littlefield. J W. McGovern. A P & Maraeson. ’ K B, Proc natl acad sci US 49 (1963) 102. 17. Friedman. D L. Biochem biouhvs . _ res comm 39 (1970) 106. ’ 18. Lindsay, J G, Berryman, S & Adams, R L P, Biochem j 119 (1970) 839. 19. Madreiter, H, Kaden, P & Mittermayer, C, Eur j biochem 18 (1971) 369. 20. Howell, S H & Hecht, N B, Biochim biophys acta 240 (1971) 343. 21. Stockdale, F E, Dev biol 21 (1970) 462. 77 Loeb. L A. Aaarwal. S S & Woodside. A M. --. ’ ’ Proc natl acad &ci US.61 (1968) 827. 23. Loeb, LA & Agarwal, S S, Exptl cell res 66 (1971) 299. 24. Bollum, F J, Cancer res 19 (1959) 561. Received April 28, 1972 Exptl CelI Res 75 (1972)
Experimental Cell Research 75 (1972) 433-441
SEA URCHIN
NUCLEAR
IV. Reversible Association
DNA POLYMERASE of DNA Polymerase with
Nuclei During the Cell Cycle’ B. FANSLER and L. A. LOEB Institute for Cancer Research, Fox Chase, Philadet’phia, Pa 19111, USA
SUMMARY We have taken advantage of the natural synchrony of sea urchin embryos during early development to study the relationship between DNA polymerase activity and nuclear DNA replication. Nuclei were isolated from early embryos (2-16 cells) in various phases of the division cell cycles. The polymerase activity found in nuclei isolated from S phase cells was 5 to lo-fold greater than that in similar fractions isolated from mitotic or G 2 embryos. As DNA synthesis is completed, the polymerase activity drops to a level slightly higher than the comparable stage before S. The ratio of DNA polymerase activity to in vivo synthesized DNA is consistently highest at the beginning of S. If mitosis is prolonged, by the isotope effect of D,O on mitotic spindle proteins, there is a corresponding delay in the increase of nuclear polymerase activity and DNA replication. We interpret these results to mean that DNA polymerase attaches to the chromosomes as they complete mitosis, functions in DNA synthesis, dissociates from them and leaves the nucleus when DNA replication is completed. This translocation is repeated with each cell cycle.
Early development of sea urchin embryos is characterized by exponential cell division with accompanying DNA replication. Our previous studies [l, 21 have shown a high level of DNA polymerase activity in unfertilized eggs and this amount of activity per embryo does not change during early development. In further studies we made a direct comparison between the DNA polymerase activity of isolated nuclei and the sea urchin embryos from which they were derived. Using embryos taken at various times between fertilization and blastula (18 h at 15”(Z), we isolated nuclei and a cytoplasmic fraction and prepared a whole embryo homogenate. DNA polymerase activity was assayed on each of these preparations. As development proceeds 1 Paper III in this series is Ref. 3. 29 -
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the activity in the nuclear fraction increases, that of the cytoplasmic decreases and the whole embryo activity remains constant. By the blastula stage, when cleavage has slowed considerably, the majority of the DNA polymerase activity is associated with the nuclei. Studies in which DNA polymerase was purified to homogeneity from embryos briefly exposed to radioactive amino acids [3] indicate that the synthesis or turnover of this enzyme during early development is no greater than most other proteins. These studies were supported by the work of Fry & Gross [4] who showed that only about 20% of the sea urchin embryo’s total protein undergoes turnover in 24 h after fertilization. Since little polymerase is synthesized during early development, we can presume that the progressive increase in nuclear polymerase activExptl Cell Res 7.5(1972)
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itv renresents a molecular transfer of the enzyme from the cytoplasm to the nucleus. However, we do not know when during the cell cycle the enzyme becomes associated with the nucleus or what happens to it after that. We now report on experiments in which nuclei were isolated from synchronously dividing embryos at several points during the cell cycle. The nuclear fractions and chromatin isolated from them were assayed for DNA polymerase activity. The results suggest that DNA polymerase attaches to the chromosomes as mitosis is completed, and then dissociates from the chromosomes. -d
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MATERIALS The sea urchins, Strongylocentrotus purpuratus were purchased from Pacific Bio-Marine, Venice, Calif. Homogeneous sea urchin DNA polymerase was prepared from nuclei of S. purpuratus blastulae (fraction VIII) [S]. Unlabeled deoxynucleoside triphosphates were purchased from CalBiochem. c@P-thymidine triphosphate was purchased from International Chemical and Nuclear Coruoration. Citv of Industrv. Calif. Calf thymus DGA was &qui;ed from Woi: thington Biochemical Corp. Deuterium water (D,O) waspurchased from Sigma Chemical Co. ‘Max&n& activated’ calf thymus DNA was prepared as described by Loeb [5]. Artificial sea water was made from distilled water and salts to the following concentration: 0.446 M NaCl, 0.0255 M MgCl,, 0.5182 M MgSO,, 0.009 M KCI. 0.0022 M NaHCO,, and 0.097 M CaCl,. This was us&d for all experim&ts unless otherwise indicated.
METHODS Preparation of hatching enzyme Sea urchin embryos were grown to the hatching stages, about 18 h at 15°C. When 95 % had visibly lost their fertilization membranes the culture was cooled to 4°C and the embryos filtered from the sea water containing the hatching enzyme. This can be kept chilled uv to 6 h without loss of activitv. Alternativelv. after filtration (Millipore, 0.45 pm), _the solution can be stored frozen at -20°C for several months. It is necessary to concentrate the thawed solution about 5 times by Diaflo ultrafiltration, UM-1 filter, as enzyme activity is lost upon freezing.
Protocol for synchrony experiments Sea urchin eggs were shed by injecting0.5 M KCl. The eggs were collected into sea water, the sperm were Exptl Cell Res 75 (1972)
collected ‘dry’ into weighing boats. Washing and fertilizing the eggs were as described by Mazia et al. 161.Eggs from one urchin (610 ml packed; about 1 in 50 females yield enough eggs for these experiments) were fertilized in 100-200 ml sea water at 15°C. Only batches in which 98 % of the eggsfertilizedwithin2min were used in these experime&& After 5 min, 6-8 vol of sea water containing hatching enzyme were added. Membranes were not visible 1 to 2 h after adding the enzyme. At a time between 50 and 130 min after fertilization, SH-thymidine (3H-TdR, spec. act. 6.7 Ci/mM) was added to the culture. The embryos were then allowed to develop to the desired stage. The precision of the synchrony was monitored at frequent intervals by microscopic examination. The timing of DNA synthesis during the cell cycle was monitored by measuring the incorporation of 3H-TdR into whole embryos. 1.0 ml aliquots, in duplicate, are removed from the developing culture (about lo* embryos) and the amount of 3H-TdR incorporated was determined by the method of Hinegardner et al. [7].
Nuclear isolation Aliquots for nuclear isolation (l-2 x 106) embryos were immediately mixed with 2 vol of a sea water ice slush to stop development. All procedures were performed at 04°C and all centrifugations are 2 500 g for 10 min unless otherwise stated. The embryos were centrifuged from the iced sea water, put into 30 ml Corex round bottom tubes and washed twice in 25 ml of 1.0 M dextrose and once in 25 ml of a hypotonic solution of 0.15 M sodium chloride with 0.015 M sodium citrate. The pellet was suspended in 12 ml of the hypotonic solution and was then dispersed by forcing the suspension 3 times through -a syringe number 20 needle. An equal volume of 2.0 M sucrose was thoroughly mixed- with the suspension after which it was centrifuged at 14 500 g for 30 min. Care was taken to decant the supernatant without contaminating the pellet with the floating yolk material. After draining the tubes and removing excess sucrose solution, the nuclear pellets were dispersed in 0.5 ml of solution E: 20 % glycerol, 0.02 M potassium phosphate buffer, pH 7.8, 4.0 mM reduced glutathione and 0.4 mM potassium versenate and sonicated intermittently at O-1°C. This is referred to as the nuclear fraction. When this method was used on cells in mitosis with no apparent nuclear membrane, the recovery of nuclear material as judged by microscopic examination and recovery of radioactive DNA in the nuclear fraction was not significantly different from nuclei in other phases of the cell cycle.
Chromatin preparation Chromatin isolation from nuclei was carried out in 20 % glycerol since polymerase requires polyglycols for stability [5]. The nuclei were suspended in 20 % glycerol, broken by intermittent sonication at 0-1°C and dialysed 18 h against 2 1of 20 % glycerol. Particulate material was removed by centrifugation at 2 500 g. The supernatant was then centrifuged at 100 000 g in an SW 40.1 rotor for 90 min. The resulting
Sea urchin nuclear DNA polymerase. IV pellets were suspended by homogenization in 1.0 ml of solution E. The nm ratio 280/260 is about 0.8.
DNA polymerase activity The assay measures the incorporation of [a+P]thymidine triohosohate into an acid insoluble nroduct. The reaction mixture, in a total volume of-O.3 ml, contains 20 % glycerol, 25 pmoles Tris-maleate buffer, pH 8.0; 4.5 pmoles MgC12; 0.3 pmole 2-mercaptoethanol; 10 nmoles each of dATP, dCTP, dGTP and [@2p]dTTP (about 3 x 10” dpm/nmole); 177 nmoles of ‘activated’ calf thymus DNA (expressed as mononucleotide and O.Oj ml of nuciea; fraction). After incubation 0.1 ml of 1.0 mg/ml denatured DNA was added and the reaction was stopped with 2.0 ml of cold 0.5 N perchloric acid containing 0.01 M sodium pyrophosphate. After standing for 10 min at O”C, the precipitate was pelleted by centrifugation at 5 000 g for 20 min. The pellets were dissolved in 0.5 ml of 0.2 M sodium hydroxide and precipitated with 1.Oml of the cold perchlorate-sodium pyrophosphate solution. After 10 min at O”C, 3.0 ml of cold water were added and the precipitate collected on a glass fiber filter (Whatman GF/C, 2.4 cm). The filter was washed once with 0.5 ml of the uerchlorate-sodium pyrophosphate solution, twice with 10 ml cold water, once with 2.0 ml of 95 % ethanol, transferred to a glass vial, dried and the radioactivity measured [2]. Protein was determined by the method of Lowry et al. [8] using crystalline pancreatic ribonuclease as g standard. Protein fractions containing interfering substances such as glutathione, 2-mercaptoethanol, MgCI, and Tris-maleate were first isolated by precipitation with cold perchloric acid.
The protocol for these experiments was to fertilize the eggs from one urchin and allow them to develop synchronously in the presence of 3H-TdR. The timing of DNA synthesis during the cell cycle was monitored by frequently taking small aliquots of embryos to determine 3H-TdR incorporation into whole embryos. At intervals of 5 or 10 min during one cell cycle, larger aliquots of the culture were taken for nuclear isolation. These nuclei, or crude chromatin isolated from them, were assayed for DNA polymerase activity using a-32P-TdRtriphosphate as the labelled substrate. The 3H-DNA which is in the nuclear fractions used as the source of polymerase also precipitates in the assay procedure. So the ratio of polymerase activ-
1.4
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-- 0.12 4--- 0.08 --0.04 e--- 0
0.6
-04
90
RESULTS
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120
150
180
210
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Fig. I. Abscissa: time after fertilization (min); ordinate: (left) O-O, aH-TdR incorporation; (right) O---O, DNA polymerase activity (cpm 52P)/thymidine incor-
poration@pm$H).
Ratio of DNA polymerase activity (““P cpm) to thymidine incorporation (3H cpm) in nuclei isolated from embryos during three cell cycles. In expts 1, 2 and 3, 10, 6 and 10 ml of eggs respectively were fertilized and the fertilization membranes removed with hatching enzyme as described in Methods. In expt 1, 0.1 mCi of 3H-TdR was added 50 min after fertilization. In expt 2, 0.3 mCi SH-TdR was added 130 min after fertilization. In expt 3, 0.2 mCi SH-TdR was added 110 min after fertilization. Nuclear isolation, polymerase assays and 3H-TdR incorporation into whole embryos were determined as in Methods. In all experiments, polymerase assays were incubated 30 min at 37°C.
ity, i.e. 32P,to in vivo synthesized DNA, i.e. 3H, can be obtained directly on the same sample. This ratio is very reliable since it is not dependent upon variations in recovery of nuclear material at different points across the Exptl Cell Res 75 (1972)
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OL Fig. 2. Abscissa: time after fertilization (min); dinate:
(left)
O-O, 3H-TdR incorporation (cpm x 1O-5); O---O nmoles TM a2P/mg protein. DNA polymerase activity per mg protein in nuclei isolated from embryos during the 4-8 cell cycle. Details are given in the caption to fig. 1 as expt 2. Synchrony refers to the incorporation of aH-TdR into DNA by the whole embryo.
(right)
cell cycle. A sharp increase in the ratio is significant as an indication that DNA polymerase has become associated with the nuclei. Fig. 1 shows the ratio of DNA polymerase activity to thymidine incorporation from three experiments over three cell cycles from 2 to 16 cells. The synchrony curves (thymidine incorporation) show that the cells were well synchronized in each experiment; having a cell cycle of 60 min with DNA synthesis occupying at most 20 min. (There is no G 1 in these cells. DNA synthesis begins in midtelophase and ends in early G2). During mitosis (140, 200, 260 min) the ratio is very low. However, with the beginning of DNA synthesis in each cycle (150, 210, 270 min) the ratio increases several-fold to its highest value. In experiment 3, nuclei were isolated over two S periods to show that this pattern of change in the ratio is consistent from one cell cycle to the next. These results indicate that the DNA polymerase becomes associated with the nucleus at the very beginning of the DNA synthetic period. As DNA synExptl Cell Res 75 (1972)
thesis is completed, the ratio decreases (160, 170, 220, 230 and 280 min), but it is difficult to determine from these data whether it was due to the increase in thymidine incorporation (tritium counts) alone, or this increase in tritium counts coupled with an actual decrease in polymerase activity. In other words, does the DNA polymerase remain attached to the chromosomes through mitosis or does it dissociate from them at the end of S? Fig. 2 gives the DNA polymerase activity of nuclei isolated from embryos at various phases of a 4-8 cell cycle. This is the specific activity data (polymerase activity per mg protein) of expt 2 in fig. 1. In contrast to the total embryo DNA polymerase activity which varies only about one-fold during the cell cycle [9], activity in the isolated nuclei increases ten-fold as the cells begin DNA synthesis. In this, and the other 4-8 cell experiment, the highest polymerase activity is in nuclei from mid-S, but in most other experiments it has been at the beginning of S. In all the experiments, however, the nuclear DNA polymerase activity decreases sharply as the cells complete DNA synthesis, reaching a minimum very soon thereafter. The most direct interpretation of these results is that DNA polymerase leaves the nucleus as replication is completed. It is to be noted that in all the experiments the polymerase activity after DNA synthesis is slightly higher than at the comparable stage before. It is possible that the increase in specific activity of polymerase in nuclei at the start of replication is simply unbound enzyme selectively trapped in the nucleus by the reformation of the nuclear membrane which occurs in early telophase. This is improbable because the nuclear membrane reforms out from the chromosomes in a manner which seemingly excludes most of the cell cytoplasm [lo]. Nevertheless, to show that polymerase is associated with chromatin, a crude fraction
Sea urchin nuclear DNA polymerase. IV was preared from the nuclei isolated over a 4-8 cell cycle. To insure the stability of the polymerase, the nuclei were suspended in 20 % glycerol and broken by sonication. After dialyzing and centrifuging out particulate matter, the chromatin was pelleted from the 20% glycerol. If this pellet is resuspended by homogenization in 20% glycerol and repelleted, there is only 5 % loss of polymerase activity. We assumed from this that the enzyme which is associated with the chromatin pellet must actually be bound to the DNA. The results of polymerase assays in these chromatin fractions are presented in fig. 3. The polymerase activity per mg protein and the ratio of polymerase activity to thymidine incorporation is low during mitosis and reach a maximum at the very beginning of S, 210 min, as in the isolated nuclei. When the DNA polymerase assays were done on these chromatin fractions without any added primer DNA, the pattern of polymerase activity over the cell cycle was the same with only about 10 Y0 of the enzyme activity. The polymerase activity also is highest at the start of DNA replication. But apart from this, the results are somewhat difficult to interpret because the amount of enzyme activity declines precipitously during DNA synthesis and actually reaches a minimum slightly before it is completed. This may be explained to some degree by the sonication used to break the nuclei; for example, the complex of chromatin and DNA polymerase may be particularly delicate and susceptible to shock during S phase. The conclusions from these chromatin assays must be drawn with some reservations. Nevertheless, because the pattern of polymerase activity over the cell cycle closely resembles that of the nuclear fractions, and because the highest ratio of polymerase activity to thymidine incorporation is at 210 min, the very beginning of S, it seems reasonable to assume that at least part of the DNA
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Fig. 3. Abscissa: time after fertilization (min); ordinate: (left) O-O, SH-TdR incorporation (cpm x 10m4); A--A (right) (outer) DNA polymerase activity (cpm TM 32Pincorporation)/thymidine incorporation (cpm 3H); (inner) nmoles TM 82P/mg protein. DNA polymerase activity in chromatin prepared from nuclei isolated at different times during the 4-8 cell cvcle. Nine ml of eaas were fertilized and the fertilization membranes k-moved with hatching enzvme as described in Methods. 0.2 mCi 3H-TdR was added at 100 min. Nuclear isolation, chromatin isolation, polymerase assays and 3H-TdR incorporation into whole embryos were determined as in Methods. Polymerase assays were incubated for 1 h at 37°C.
polymerase entering the nucleus at the beginning of DNA synthesis becomes bound to the chromosomes. Is this decrease in polymerase activity at the end of S actually due to a loss of enzyme, or possibly some inhibitor which appears at that time? In order to check for enzyme inhibition, purified sea urchin DNA polymerase was mixed with nuclei having the lowest activity and we looked for additivity of the enzyme activities. For example, in an experiment in which the activity for the nuclear fraction (time 250, expt 3, fig. 1) was 250 pmoles TMP incorporated, and the purified enzyme incorporated 120 pmoles, together the activity was 390 pmoles. This indicates additivity between the purified and nuclear enzyme activities thus ruling out any readily soluble inhibitor present in excess under the conditions of the in vitro assay. Exptl Cell Res 75 (1972)
438 B. Fader & L. A. Loeb
Fig. 4. Abscissa: time after fertilization (min); ordinate: (left) 3H-TdR incorporation (cpm x 10--4). O-O, D,O treated embryos; O-O, untreated embryos; (right) o---n, nmoles TM 52P/mg protein. Effect of D,6 on DNA synthesis and nuclear DNA polymerase activity. Seven ml of eggs were fertilized and thefertilization membranes removed with hatching enzyme as in Methods. After 90 min the embryos were allowed to settle and the sea water volume reduced to 125 ml. 0.1 mCi of SH-TdR was added at 130 min. A 10 ml portion of the culture was removed and allowed to develop normally. From this, 0.5 ml synchrony aliquots were taken at times indicated in order to monitor normal DNA synthesis. At 180 min after fertilization 15 ml of the culture were taken for nuclear isolation. Immediately, 100 ml of D,O sea water were added to the remaining 100 ml of the culture to give a 50 % D,O final concentration. One ml synchrony aliquots were taken from time 180 to time 200. At 190 and 210 min, 30 ml aliquots were taken for nuclear isolation. Immediately after the 210 min aliquot was taken, 1 440 ml sea water were added to give a final D,O concentration of 5 % which allows the embryos to develop normally. After this time, 10 ml synchrony and 300 ml nuclear isolation aliquots were taken. All procedures for nuclear isolation and polymerase assays are as in Methods. Assays were incubated 30 min at 37°C.
We then wanted to know if this pattern of increase and decrease in DNA polymerase activity over the cell cycle is unique to this enzyme, or might it be common to other enzyme activities under these conditions. To answer this, nuclear fractions were assayed for three other enzyme activities, DNase, thymidine kinase, and lactic acid dehydrogenase; chosen because of their widely diverse functions with respect to cellular metabolism. While there are variations in these enzyme Exptl Cell Res 75 (1972)
activities over the cell cycle, none of the three in any way shows the consistent pattern of change which is observed for nuclear DNA polymerase activity. It appears, then, from these studies that the increase and decrease of nuclear DNA polymerase activity with the onset and completion of each S period is a property of this enzyme. To show a correlation between the rise in polymerase activity and the beginning of DNA synthesis, synchronized embryos were reversibly stopped in metaphase. The delay was accomplished by adding an equal volume of 100% deuterium oxide (DzO) sea water to a culture of embryos in prophase. Gross & Spindel [11] have shown that echinoderm embryos in 50% D,O sea water will not develop past metaphase, presumably because of an isotope effect preventing the mitotic spindle subunits from dissociating. This inhibition is reversed within 1 min of lowering the D,O concentration to 5 %. Embryos were in 50 Y0 D,O for a 30 min period from 180 to 210 min post-fertilization and then diluted into 9 vol of sea water without DzO. Aliquots were taken and nuclei isolated from them at the points indicated in fig. 4. In addition, a small amount of the culture not exposed to D,O was allowed to develop in order to monitor normal DNA synthesis. Fig. 4 shows the DNA polymerase activity of nuclei isolated from cells during and after D,O inhibition. DNA synthesis is delayed about 25 min, from time 220 to about 245 min after fertilization. Nuclear DNA polymerase activity decreases through the period of D,O treatment and after, until, at mid-telophase it shows the characteristic marked increase. We conclude that when mitosis is extended, there is a proportional delay in the association of DNA polymerase with the chromosomes and in the subsequent DNA synthesis.
Sea urchin nuclear DNA polymerase. IV DISCUSSION We have presented evidence which suggests that the DNA polymerase in early developing sea urchin embryos becomes associated with the nucleus at the very beginning of DNA synthesis and subsequently leaves the nucleus as replication is completed. Furthermore, at least part of the nuclear polymerase attaches to the chromatin during S, although the association may be rather delicate and easily disrupted. This pattern of reversible association is repeated in each cell cycle with the DNA polymerase presumably residing in the cytoplasm during G2 and mitosis. The decrease in polymerase activity at the end of S is not due to a soluble inhibitor present in excess. If mitosis is prolonged by the isotope effect of D,O on mitotic spindle proteins, there is a corresponding delay in the increase of nuclear DNA polymerase activity and DNA replication. These results should be considered in the context of the nucleo-cytoplasmic events during the cell cycle. At the second, third and fourth cleavages only 5-15 % of the unfractionated embryo’s total DNA polymerase activity can be recovered in isolated nuclei. This enzyme activity and what seems more significant, the activity associated with chromatin shows an abrupt several-fold increase at the beginning of each S. In the sea urchin embryo this is the point in mitosis when the chromosomes begin to extend after being in intimate contact with the cytoplasm. Presumably, as the chromosomes unwind, many points on the DNA become available for enzyme attachment. Loeb [12] has estimated, on the basis of the polymerase activity bound to a crude chromatin preparation, from the amount of activity of the homogeneous enzyme, and the amount of DNA per embryo, that one DNA polymerase molecule could bind for each two to three thousand nucleo-
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tide pairs. The binding points on the DNA are presumably 3’-OH groups. Loeb [5] has shown that DNA treated with pancreatic DNase to produce a 3’-OH is an excellent primer for sea urchin polymerase while DNA digested with micrococcal nuclease to give 3’-PO, will not prime. These binding points, could result from the activity of an endonuclease similar to that described by Burgoyne et al. [13]. They reported a calciumdependent endonuclease activity which enables DNA polymerase in rat liver nuclei to use the nuclear DNA as primer-template. After DNA synthesis is completed, the DNA polymerase activity of both the nuclear and chromatin fractions decreases. Since the polymerase activity of the whole embryo remains fairly constant throughout the cell cycle, and indeed, through early development up to gastrulation, we have concluded that this decline in activity is enzyme leaving the chromosomes and the nucleus. However, the possibility of some enzyme inhibitor appearing at the end of S cannot be completely excluded. This reversible association of DNA polymerase with the nucleus during the mitotic cycle was unexpected since it creates an apparent paradox with the results reported earlier from this laboratory [2], that polymerase activity accumulates in the nucleus as cleavage progresses. Fig. 2 shows that polymerase activity after DNA synthesis is slightly higher than at the comparable stage before S. We found this in all the experiments and interpret it to be residual DNA polymerase which remains with the chromosomes when they condense at prophase. This residual activity could accumulate with each cell division during early development and is sufficient to account for the translocation of polymerase activity into nuclei which was observed in the previous study [2]. These results correlate well with Mazia’s Exptl Cell Res 75 (1972)
440 B. Fader & L. A. Loeb [14] observations. In different partially synchronized populations, he counted the number of sea urchin embryos which were in mitosis and measured the DNA polymerase activity associated with a crude chromatin fraction prepared from them. The polymerase activity was highest in chromatin from cells which had just finished mitosis. He proposed that the enzyme attached to the chromosomes at the end of mitosis, replicated the DNA and then dissociated from them before the next mitosis. Other studies [15, 161 on cells which were synchronized by chemically blocking the DNA synthesis with 5-fluoro-2’deoxyuridine showed a small redistribution of DNA polymerase activity from cytoplasmic to nuclear fraction upon release of the block. Friedman [ 171, using HeLa cells, synchronized them by prolonged exposure to amethopterin and adenosine to induce a thymidineless condition. He then induced DNA synthesis by the addition of thymidine and found that nuclear DNA polymerase activity increased as the cells went into S and decreased as it was finished. Lindsay et al. [18] could not demonstrate the same effect in L cells, synchronized with aminopterin. Madreiter et al. [19], also using L cells, synchronized them mechanically in order to avoid the physiological imbalance of blocking agents. They showed a rise in the level of DNA polymerase activity in a particulate fraction as DNA synthesis occurred. They also looked at DNase activity associated with the same fraction and found little variation over the cell cycle. Howell & Hecht [20] have reported on the DNA polymerase and polynucleotide ligase activities during the mitotic cell cycle of the naturally synchronous LiEium microspore cells. They looked only at the whole cell supernatant fraction after high speed centrifugation. There was no significant fluctuation in DNA polymerase activity during the cell cycle, however, there was a sharp Exptl Cell Res 75 (1972)
increase in polynucleotide ligase activity before S which then decreased during S and subsequently disappeared. They suggest that in this system, the amount of ligase activity may have some controlling influence in DNA synthesis. Each of these studies has the disadvantage of either having to establish synchrony by chemical blocks, the total effects of which are unknown, or mechanically choosing cells in one phase of the cell cycle and having only partial synchrony. In the case of Lilium microspores, the study unfortunately included only enzyme activity in whole cell supernatants. It therefore becomes difficult to compare them directly with our studies, but generally they suggest that nuclear DNA polymerase activity is high as DNA is replicated. Friedman’s work further suggests that the enzyme leaves the nucleus after S. None of them, however, establishes any real correlations between naturally induced DNA synthesis and the level of nuclear DNA polymerase activity. With respect to the control of DNA synthesis, the early developing embryo represents an entirely different situation from a fully differentiated eucaryotic cell. The embryonic cells are dividing rapidly and need a large amount of enzyme to replicate their DNA. On the other hand, differentiated cells generally function in their individual ways, in most tissues they seldom divide and do not require large amounts of DNA polymerase. There is evidence which indicates that in some systems the presence or absence of polymerase activity may be one of the primary factors controlling replication. Stockdale 1211 has shown a very close correlation between the decline of DNA polymerase activity and the lack of ability of multinucleated muscle cells to synthesize DNA or divide. He demonstrated a one hundred-fold decrease in the amount of enzyme activity as the mononucleated cells fused to form the multi-
Sea urchin nuclear DNA polymerase. IV nucleated non-dividing skeletal muscle cells. He was not able to determine if the sharp decrease was due to a loss of the enzyme or an accelerated inactivation. Conversely, when lymphocytes are stimulated with phytohemagglutinin there is a long delay before the cells divide. Loeb et al. [22] have shown that during this delay the cells must synthesize DNA polymerase, after which there is then an excellent correlation between the amount of polymerase activity and the rate of DNA synthesis [23]. There are systems, however, where DNA synthesis does not seem to be dependent upon the level of DNA polymerase; for example, in regenerating rat liver DNA synthesis ceases, but the level of DNA polymerase activity remains high [24], indicating that the cessation of DNA synthesis in this system does not depend upon the loss of enzyme activity. The control of DNA synthesis in the sea urchin embryo does not depend upon the synthesis of new DNA polymerase. There is a high level of activity in the unfertilized egg which persists through the blastula stage. Instead, the control mechanism may involve some step in the transfer process of the enzyme from the cytoplasm to the nucleus; possibly the availability of enzyme attachment sites on the chromosome. We can now imagine a series of events as follows. A molecule of DNA polymerase, which has been synthesized during oogenesis and stored in the cytoplasm, comes in contact with a chromosome unwinding from its mitotic configuration. The enzyme attaches, presumably to a 3’-OH DNA terminus which could result from the action of an appropriate endonuclease. The polymerase participates in DNA replication, dissociates from the chromosome and probably leaves the nucleus altogether. A residual polymerase activity remains associated with the chromosomes through mitosis and accmulates in the nucleus with each cell division. This translocation is
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repeated with each cell cycle, as enough enzyme is made during oogenesis in the sea urchin to accommodate the rapid DNA synthesis through the blastula stage. We thank Dr Jack Schultz for generous counsel. and dedicate this paper to his memory. This investigation was suooorted by USPHS arants CA-l 1524 and CA-l 1835 from NIH and gram GB18419 from NSF. Support was also derived from grants to this Institute: LJSPHS, CA-06927 and RR05539, and an appropriation from the Commonwealth of Pennsylvania.
REFERENCES 1. Loeb, L A, Fansler B, Williams R & Mazia, D, Exptl cell res 57 (1969) 298. 2. Fansler, B & Loeb, LA, Exptl cell res 57 (1969) 305. 3. Loeb, L A & Fansler. B. Biochim bioohvs . < acta ’ ’ 217 (1970) 50. 4. Fry, B J & Gross P R. Dev biol21 (1970) 125. 5. Loeb L A, J biol them 244 (1969) l672.’ 6. Mazia, D, Mitchison, J M, Medina, H & Harris, P J, J biophys biochem cytol 10 (1961) 467. 7. Hinegardner. R T. Rao. B & Feldman. D E. Expti cell res 36 (1964) 53. 8. Lowry, 0 H, Rosebrough, N J, Farr, A L & Randall, R J, J biol them 193 (1951) 265. 9. Loeb, L A, Fansler, B & Salter J P, Symposium on uptake of informative molecules by living cells (ed L Ledoux). North-Holland, Amsterdam and London (1970). 10. Mazia D, The cel1 (ed J Brachet & A E Mirsky) vol. 3, ^_ DU. 77-412. Academic Press. New York and London (1961). Il. Gross, P R & Spindel, W, Ann NY acad sci 90 (1960) 500. 12. Loeb, L A, Nature 226 (1970) 448. 13. Burgoyne, .L A, Waqar, M A‘& Atkinson, M R, Biochem biophys res comm 39 (1970) 254. 14. Mazia, D, Cell nucleus, pp. 15-25. Taylor & Francis, London (1966). 15. Gold, M & Helleiner, C W, Biochim biophys acta 80 (1964) 193. 16. Littlefield. J W. McGovern. A P & Maraeson. ’ K B, Proc natl acad sci US 49 (1963) 102. 17. Friedman. D L. Biochem biouhvs . _ res comm 39 (1970) 106. ’ 18. Lindsay, J G, Berryman, S & Adams, R L P, Biochem j 119 (1970) 839. 19. Madreiter, H, Kaden, P & Mittermayer, C, Eur j biochem 18 (1971) 369. 20. Howell, S H & Hecht, N B, Biochim biophys acta 240 (1971) 343. 21. Stockdale, F E, Dev biol 21 (1970) 462. 77 Loeb. L A. Aaarwal. S S & Woodside. A M. --. ’ ’ Proc natl acad &ci US.61 (1968) 827. 23. Loeb, LA & Agarwal, S S, Exptl cell res 66 (1971) 299. 24. Bollum, F J, Cancer res 19 (1959) 561. Received April 28, 1972 Exptl CelI Res 75 (1972)