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DEVELOPMENTAL
BIOLOGY
79,
119-127
Limited Complexity
(1980)
of the RNA in Micromeres
Sea Urchin Embryos
of Sixteen-Ceil
SUSAN G. ERNST,’ BARBARA R. HOUGH-EVANS, ROY J. BRITTEN,~ AND
ERIC H. DAVIDSON
Division of Biology, California
Institute of Technology, Pasadena, California 91125 and Kerckhoff Marine
Lab of the Division of Biology, California
Znstitute of Technology, Corona de1 Mar, California
92625
Received April 2, 1980; accepted April
11, 1980
The sequence complexity of sea urchin embryo micromere RNA is about 75% of that of total
16-cell embryo cytoplasmic RNA, as reported earlier by Rodgers and Gross [Rodgers, W. H., and
Gross, P. R. (1978) Cell 14, 279-2881. In contrast to the rest of the embryo, there are few, if any,
complex maternal RNA species in the micromere cytoplasm which are not represented in the
polysomes. The micromeres do not contain detectable quantities of high-complexity
nuclear RNA,
though such RNA exists in other cells of the fourth-cleavage
embryo.
INTRODUCTION
thus give rise to a discrete cell lineage, the
differentiated
character of which is well
defined in morphological terms.
The origin of the functional distinction
between the micromere cell lineage and the
remainder of the embryo is a matter of
speculation. Among the possibilities often
considered are that the 1Bcell embryo contains an unequal distribution
of maternal
mRNA sequences or of newly synthesized
embryo mRNAs. However, Senger and
Gross (1978) reported that protein synthesis patterns in the three blastomere cell
types are qualitatively
similar. Even with
the increased resolution made possible by
two-dimensional
gel electrophoresis,
Tufaro and Brandhorst
(1979) detected no
clear qualitative
differences among the
three cell types in the synthesis of more
than 1000 different proteins. Thus it would
appear that micromeres contain more or
less the same set of abundant and moderately abundant mRNAs as does the rest of
the 16-cell embryo. On the other hand,
Rodgers and Gross (1978) found that the
rare single-copy sequence transcripts of the
16-cell embryo are not homogeneously distributed. Their experiments indicated that
micromere RNA includes only 6580% of
At the eight-cell stage of early cleavage,
sea urchin embryos consist of four “animal”
blastomeres and four “vegetal”
blastomeres, all of which are similar in size and
form. During fourth cleavage, the animal
blastomeres again divide equally, giving rise
to eight mesomeres, but the four vegetal
blastomeres cleave unequally
into four
large macromeres and four very small micromeres situated at the vegetal pole of the
embryo. All of the blastomeres, including
the micromeres, undergo several further
rounds of division as the embryo assumes
the hollow blastula form. Descendants of
the micromeres, the primary mesenchyme
cells, then migrate into the blastocoel.
Groups of these cells secrete two calcareous
granules, upon which the first spicules of
the larval skeleton are later constructed
(Horstadius, 1937, 1973). Micromeres that
are isolated and maintained in culture appear to follow their normal developmental
fate, including the secretion of skeletal spicule elements (Okazaki, 1975). Micromeres
’ Present
address: Department
of Biology,
Tufts
University, Medford, Massachusetts 02155.
’ Also staff member, Carnegie Institution
ington.
of Wash119
0012-1606/80/110119-09$02.00/O
Copyright 0 1980 by Academic Press, Inc.
All rights of reproduction in any form reserved.
120
DEVELOPMENTAL
BIOLOGY VOLUME79,198O
the complex set of low-abundance maternal
RNA sequences found in the remainder of
the embryo.
In this report we present further measurements of the complexity of micromere
total RNA and, in addition, of micromere
polysomal RNA. The results imply that, in
molecular terms, micromeres differ from
the remainder of the embryo in several
ways that may be relevant to their separation as a unique cell lineage.
MATERIALS
AND METHODS
Sea urchin embryos. Strongylocentrotus
purpuratus females were spawned and the
eggs washed, collected, and fertilized by
standard methods (Smith et al., 1974). Fertilization was at a concentration of about 25 x 105 eggs/ml Pen-Strep MPFSW (30
units/ml penicillin,’ 50 pg/ml streptomycin
in Millipore-filtered seawater). The fertilized eggs were demembranated by modifications of standard procedures (Hynes and
Gross, 1970). At 90 set or when 85-90%
fertilization was achieved, whichever came
first, an equal volume of freshly prepared
0.08% papain, 0.40% glutathione (pH 7.8)
was added. The eggs were kept in suspension by gentle swirling, and fertilization
membranes began to disappear within 90120 sec. After 7-9 min, the eggswere diluted
to l-3 x 104/ml and grown at 14-15’C with
stirring. Demembranization by this method
did not affect the normal development of
the embryos to the pluteus stage.
In some experiments RNA was isolated
from whole 16-cell embryos. These embryos
were demembranated, grown, and dissociated as described for isolation of micromeres. Thirty-two- to 64-cell embryos were
cultured by standard procedures (Smith et
al., 1974) and harvested at 7.5 hr, when
approximately 50% of the embryos had undergone the sixth division and contained 64
cells, while the remainder were still at the
fifth-cleavage stage.
Isolation of micromeres. Micromeres
were isolated by a procedure similar to that
employed by Rodgers and Gross (1978).
About 5 hr after fertilization, when the
embryos had just completed their fourth
division, they were allowed to settle and
the seawater was aspirated off. The growing
chamber was placed in an ice bath to retard
further cell division and “run off’ of ribosomes from polysomes. Eight hundred to
1200 ml of ice-cold CMFSW (calcium-magnesium-free seawater) was added to the
embryos to dissolve the hyalin layer and
dissociate the embryos. Two or three
washes of CMFSW were generally sufficient to cause dissociation. Occasionally,
blastomeres continued to adhere after several washes with CMFSW, and in this case
the solution was brought to 0.5-2.5 n&f
EDTA or to 1 it4 glycine (pH 8). Either
treatment results in complete dissociation
of the blastomeres.
Dissociated blastomeres were adjusted to
a concentration of about 3 x lo5 embryos/
ml of 1%Ficoll400,009 (Sigma) in CMFSW.
Three to 3.5 ml of cell suspension was layered onto 5-15% Ficoll gradients in
CMFSW in 30-ml Corex centrifuge tubes.
The gradients were centrifuged for 1 min at
200-250g in a Sorvall HB4 swinging-bucket
rotor. A discrete band of micromeres
formed about 25 mm from the top of the
gradient. Micromeres from 8-10 gradients
were pooled, concentrated, and, if necessary, sedimented into a second gradient.
The micromere band was harvested and a
sample containing several hundred blastomeres was counted to determine the purity
of the micromere fraction. If less than 99%
of the cells were micromeres, the preparation was repurified or discarded.
RNA preparations. Total RNA was extracted from mature eggs, dissociated 16cell embryos, isolated micromeres, and 32-,
64-cell embryos, according to the procedure
of Hough-Evans et al. (1977) for total egg
RNA. Briefly, eggs or embryos were homogenized in 7 M urea in low-salt buffer,
and the homogenate was extracted with
phenol and chloroform:isoamyl alcohol.
Nucleic acids were precipitated and the
RNA was further purified by DNase and
ERNST ET AL.
Micromere
proteinase K digestion and chromatography on Sephadex G-100. Cytoplasmic RNA
was prepared from 16-cell embryos by a
similar procedure, after cell lysis and pelleting of nuclei, as previously described
(Hough-Evans et al., 1977). Sixteen-cell
embryo polysomes were isolated according
to Galau et al. (1976). Micromere polysomes were obtained as follows: Isokinetic
5-20s sucrose gradients in 500 mM KCl, 10
mM MgC12,10 mM Pipes [piperazine-N- N’bis(2-ethanesulfonic acid)] (pH 6.5), 50 pg/
ml polyvinyl sulfate, were prepared in 5-ml
polyallomer tubes containing a 0.6~ml cushion of 40% sucrose in the same buffer. Seventy-five to 150 ~1of the postmitochondrial
supernatant fraction from the isolated micromeres was layered onto the gradients
and centrifuged at 50,000rpm in a Beckman
SW 50.1 rotor for 40 min at 4°C. The polysomes were sedimented to the top of the
40% sucrose cushion. Gradients were
pumped through an Isco flow cell and monitored for optical absorbance at 254 mn.
The <90 S material was discarded and the
polysome fraction was diluted three- to
fourfold. The polysomes were dissociated
by the addition of 0.5 M EDTA to a final
concentration of 75-100 mJ4 and incubation for 15 min on ice. The suspension was
then layered onto a second sucrose gradient. RNA was extracted from material
sedimenting at <80 S as described previously (Galau et aE.,1976).
Single-copy DNA and “egg DNA”
tracers. Total single-copy DNA was prepared as described by Galau et al. (1976).
The DNA was labeled in vitro with 3H by
the “gap translation” method with Escherichia coli DNA polymerase I (Boehringer
Mannheim) (Galau et al., 1976; Hough-Evans et al., 1977). The single-copy tracers
used had specific activities of 0.6-l x lo7
cpm/pg and a weight-average fragment size
of 200-250 nucleotides.
Egg DNA consists of that fraction of
single-copy DNA sequences which form duplexes with egg RNA. The egg DNA tracers
were prepared as previously described. An
121
RNA Complexity
excess of total egg RNA was reacted with
single-copy r3H]DNA (Hough-Evans et al.,
1977, 1979) to equivalent RNA Cot values
greater than 50,000. The hybridized [3H]DNA was harvested and further enriched
for sequences homologous to egg RNA by
a second reaction with egg RNA, followed
by isolation of the hybridizing sequences.
Four different egg DNA preparations were
used in this study.
Hybridization conditions. In the reactions described, the RNAs were in 103- to
104-fold mass excess with respect to [3H]DNA tracers. Hybridization reactions were
carried out in 0.41 M phosphate buffer (pH
6.8), 1.5 mM EDTA, 0.05% sodium dodecyl
sulfate at 60°C. All Cot values presented
are equivalent COt’s, i.e., they have been
corrected for acceleration in hybridization
rate due to Na+ concentration greater than
0.18 M (Britten et al., 1974). DNA-DNA
and DNA-RNA duplexes were assayed by
hydroxyapatite
chromatography, essentially according to Galau et al. (1974). After
hybridization, samples were diluted in 1.0
ml of 0.05 M phosphate buffer and divided
into two aliquots. One-half was brought to
0.12 M phosphate buffer and assayed for
total duplex. The other half was incubated
with 10pg/ml RNase A in 0.05 M phosphate
buffer for 1 hr at 37°C. It was then brought
to 0.12 M phosphate buffer and assayed for
DNA-DNA duplex. The difference in the
amount of bound [3H]DNA in total duplex
and in DNA-DNA duplex measures the
extent of DNA-RNA hybridization. Values
were corrected for reactivity of the [3H]DNA preparation (SS-90% for the singlecopy r3H]DNAs used). Hybridization data
were reduced by a nonlinear least-squares
procedure (Pearson et al., 1977), assuming
pseudo-first-order kinetics.
RESULTS
Micromere Preparations
Various stages in the isolation of micromeres are shown in Fig. 1. The final preparations, such as that illustrated in C, consisted of >99% micromeres. Preparations of
122
DEVELOPMENTAL BIOLXXY
at least this degree of purity were obtained
routinely by the procedures given in Materials and Methods and were used for all
the experiments described subsequently.
Complexity of Micromere RNA
Total RNA was extracted from isolated
micromeres, and its complexity was calculated from the amount of single-copy [3H]DNA tracer hybridized at apparent kinetic
termination. For comparison, reactions
were also carried out with total RNA of 16cell and 32- to 64-cell embryos. Figure 2
shows that micromere RNA hybridizes
with 2.6% of the reactable single-copy
DNA. The single-copy sequence content of
the S. purpuratus genome is 6.1 x lOa nucleotide (nt) pairs (Graham et al., 1974),
and given that single-copy sequences are
represented asymmetrically in sea urchin
RNAs (Hough et al., 1975; Lev et al., 1980),
the complexity of micromere RNA is calculated to be about 3.2 x 10’ nt. This is
slightly lower than the complexity established for the total egg RNA of S. purpuratus, 3.7 X 1O’nt (Galau et al., 1976;HoughEvans et aZ., 1977). Though a relatively
large amount of terminal data are presented
in Fig. 2 for the micromere RNA reaction,
there is no evidence for higher complexity
components. The surprising implication is
that micromeres lack detectable quantities
of the high-complexity nuclear RNA observed previously in sea urchin embryos of
later stages (Hough et al., 1975; Kleene and
Humphreys, 1977; Wold et al., 1978; Ernst
et aZ., 1979).
The hybridization experiments with 16cell and 32- to 64-cell embryo RNAs shown
in Fig. 2 demonstrate that a high-complexity micromere RNA component would easily have been noticed had it been present.
RNA extracted from dissociated but unfractionated 16-cell embryos reacts with at
least 8% of the single-copy DNA tracer.
Therefore, an RNA whose complexity is
significantly greater than that of egg RNA
exists in the macromeres and/or meso-
VOLUME 79.1980
meres of the fourth-cleavage embryo. This
high-complexity RNA is almost certainly
nuclear since previous studies have shown
that in the 16-cell embryo, as in later embryos, the cytoplasmic RNAs are of equal
or lower complexity than unfertilized egg
RNA (Hough-Evans et al., 1977). RNA
from 32- to 64-cell embryos also includes a
very complex set of sequences. The 32- to
64-cell RNA hybridization reaction proceeds more rapidly, as might be expected
since there are now more than two times as
many nuclei in the same mass of cytoplasm.
This is apparently due to an increase in the
concentration of complex sequences in the
total RNA. The dashed lines in Fig. 2 (see
legend) represent the kinetics of these reactions calculated on the assumption that
the complexity of the nuclear RNA in both
the 16-cell and the 32- to 64-cell embryos is
about the same as in later embryos, i.e.,
that the reaction terminates at 15%. However, this could not be explicitly demonstrated, since to do so would have required
reactions to be carried out to COt> 5 x lo6
M sec.
Consider the situation that would result
if each of the micromeres contained the
same amount of high-complexity nuclear
RNA as appears to be present (on the average) in the remaining cells. The four micromeres include only 7.5% of the cellular
volume of the embryo, and their nuclei
populate only about 3.3% of the total cytoplasmic volume (calculated from the dimensions of nuclei and cells in dissociated
blastomere preparations). We assume that
in micromeres the concentration of ribosomes per unit cytoplasmic volume is not
significantly greater than elsewhere in the
embryo. In this case the experiments of Fig.
2 would have revealed a high-complexity
micromere RNA reaction occurring at a
rate about 10 times faster than that observed with whole 16-cell embryo RNA. We
observe a complexity equal to about 85% of
that occurring in unfertilized egg RNA.
Since no higher-complexity reaction could
ERNST ET AL.
Micromere
RNA Complexity
123
FIG. 1. Micromere isolation. A l&cell embryo (5.5 hr after fertilization)
is shown in A. The fertilization
membrane was removed 90 set after the addition of sperm by a solution of 0.04% papain, 0.2% glutathione, pH
7.8. The hyalin layer is visible surrounding the embryo. Micromeres are indicated by the arrow (4. Embryos
were dissociated in ice-cold calcium-magnesium-free
seawater, and the resulting cell suspension is seen in B.
Arrow indicates four micromeres. There are two macromeres above and two below the micromeres. At the X,
four mesomeres can be seen above the upper two macromeres. C, taken at the same magnification,
shows a
preparation of isolated micromeres after Ficoll gradient separation. Scale bars represent 50 pm.
be detected, we conclude that there is a
clear difference between the micromeres
and the other cells of the fourth-cleavage
embryo. The micromeres could contain-if
any-no
more than a few percent of the
amount of high-complexity
RNA found per
124
DEVELOPMENTAL BIOLOGY
nucleus in the mesomeres and/or
meres.3
VOLUME
79, 1980
macro-
Maternal RNA Sequence Distribution
The purpose of the following experiments
was to determine the fraction of the egg
RNA sequence set present in micromeres
and in micromere polysomal RNA. For
these measurements, we utilized a singlecopy C3H]DNA tracer fraction (“egg DNA”)
which consists largely of sequences represented in unfertilized
egg RNA. As described by Hough-Evans et al. (1977) and
in Materials and Methods, egg DNA tracers
were prepared by two successive rounds of
hybridization
of a single-copy tracer with
egg RNA. In Fig. 3, the reactions of egg
DNA with egg RNA and with micromere
RNA are compared. Apparently only about
75% of the egg RNA sequence set is represented in the RNA of the micromeres. A
similar result was reported by Rodgers and
Gross (1978). Since the complexity of S.
purpuratus egg RNA is 3.7 x lo7 nt (Galau
et al., 1976; Hough-Evans et al., 1977), the
length of single-copy sequence shared by
both egg and micromere RNAs is about 2.7
x lo7 nt. This value is less than the total
complexity of micromere RNA measured
by reaction with single-copy DNA in the
experiment of Fig. 2, i.e., 3.2 x lo7 nt. If
real, this discrepancy implies the existence
3 Rodgers and Gross (1978) also measured a relatively low micromere RNA complexity, considering
the sum of their “egg DNA” and “null egg DNA”
reactions. However, either their total 16-cell embryo
RNA reaction with the “null egg DNA” tracer failed,
in that, for some reason, no hybrid was recovered, or
there is a species difference between Lytechinuspictus
and S. purpuratus. In any case, they concluded that
the concentration of complex nuclear RNA in the 16cell embryo is too low for any reaction to have been
observed and that micromeres probably contain such
RNA. On the other hand, the data in Fig. 2 of this
paper show that even though the 16-ceII embryo RNA
hybridizations
of Rodgers and Gross were carried out
only to Cot 5 X 104, a signifkant reaction should have
been clearly evident in their null egg DNA hybridixations. We believe that all of the measurements presented by Rodgers and Gross are consistent with ours,
except for the one negative control with total 16-cell
embryo RNA.
‘1 \
‘\ \
‘->--.
16
IO'
I
103
L
104
\
105
1
I06
RNA Cot
FIG. 2. Hybridization
of single-copy [“HIDNA with
total RNA from 32- to 64-ceil embryos, 16-cell embryos, and isolated micromeres. Total RNA was isolated from embryos at the 32- to 64-cell stage, 7.5 hr
after fertilization
(0); 16-cell embryos, 5.5 hr after
fertilization
(0); and 16-cell stage micromeres isolated
as described in Materials and Methods and shown in
Fig. 1 (A,). The RNAs were reacted in excess with
single-copy [3H]DNA tracers. The values shown have
been corrected for the reactivity (with DNA) of the
[3H]DNA tracers used (8640%). At apparent tennination, 2.6% of the C3H]DNA was hybridized by micromere RNA (solid line). The dashed lines show the
least-squares solutions to the kinetics of the 32- to 64ceII and 16-cell RNA reactions, on the assumption that
both reactions would terminate at 15%. This is the
approximate terminal value for nuclear RNA (or total
RNA) reactions observed in studies on later embryos
(Hough et al., 1975; Kleene and Humphreys,
1977;
Wold et al,, 1978; Ernst et al., 1979; unpublished data).
For the 16-cell embryo RNA, the rate constant in the
solution shown is 6.8 X lo-” M-’ set-‘; and for the 32to 64-cell embryo RNA, 1.2 x IO-” M-’ set-‘.
of nonmaternal
sequences in the micromeres. However, the difference is only 5 x
lo6 nt, close to the limit of resolution of
these procedures. Some support for the possibility that micromeres possess a special
set of single-copy RNA transcripts comes
from the experiments of Rodgers and Gross
(1978). They found that a tracer largely
depleted of egg RNA sequences (“nub egg
DNA”) reacts to about 1% with micromere
RNA. This value implies independently
that micromeres contain a nonmaternal set
of RNA sequences of about lo’-nt complexity. However, due to the small magnitude
ERNST ET AL.
B
8
Micromere
125
RNA Complexity
20
00
0
I
40
I
20
I
60
I
80
RNA Cot x l0-3
FIG. 3. Hybridization
of egg [3H]DNA with egg RNA, total micromere RNA, and polysomal micromere
RNA. The solid curve shows reactions of egg DNA tracers with excess egg RNA (e). Egg DNA is a single-copy
tracer consisting mainly of sequences complementary
to egg RNA. Data were pooled from four egg DNA
preparations by normalizing the average terminal value obtained with each preparation to 100%. These values
ranged from 52 to 60% for the various tracers. Since only 3% of the starting single-copy tracer reacts with egg
RNA, the egg RNA sequences in these preparations had been concentrated 17- to 27-fold. The pseudo-firstorder rate constant of the solution shown is 2.3 x 10m4M-’ set-‘, as reported earlier (Hough-Evans et al., 1977).
The reactions of egg DNA with total micromere RNA (A) and with polysomal micromere RNA (V) are shown
relative to the egg DNA-egg RNA reaction. The dashed line at 73% represents the reaction of a similar egg
DNA tracer with total 16-cell embryo polysomal RNA (data originally presented by Hough-Evans et al., 1977).
of all of these values, this cannot be considered a secure conclusion.
The egg DNA was also hybridized with
RNA extracted from the polysomes of isolated micromeres. These data are shown by
the open symbols in Fig. 3. This result
contrasts with the situation elsewhere in
the fourth-cleavage embryo. Thus, HoughEvans et al. (1977) showed that while the
whole maternal RNA sequence set can be
recovered from 16-cell embryo cytoplasm,
only about three-fourths of these sequences
are associated with polysomes. It follows
that the micromeres lack the nonpolysomal
cytoplasmic sequences inherited by macromeres and/or mesomeres from the unfertilized egg.
DISCUSSION
The experiments
presented here, together with those of Rodgers and Gross
(1978), show that in several respects the
population of single-copy sequence transcripts in fourth-cleavage micromeres is distinct from that in the remainder of the
embryo. A brief summary of these differences follows: (a) Micromeres lack detectable quantities of the high-complexity
nuclear RNA found in the macromeres and/
or mesomeres; (b) micromeres contain only
75% of the set of maternal RNA sequences
present in both the unfertilized egg and the
cytoplasm of the whole 16-cell embryo; and
(c) micromeres lack the nonpolysomal maternal RNA sequences found in the cytoplasmic fraction of the whole embryo. In
addition, though the evidence is not conclusive, micromeres may contain a small set of
transcripts not found in the unfertilized egg.
Such a sequence set would not have been
detected in earlier studies on the complexity of 16-cell embryo RNA (Hough-Evans
et al., 1977) because the micromeres contain too small a fraction of the embryo
cytoplasmic RNA.
The most likely explanation of the lack
of high-complexity
nuclear RNA in the 16cell micromeres is late activation of RNA
synthesis. According to De Petrocellis et al.
(1977), the first four divisions of the sea
126
DEVELOPMENTAL BIOLOGY
urchin embryo (i.e., Paracentrotus Zividus)
are synchronous, but after this the micromeres divide at a slower rate. Autoradiograph and uptake experiments show that
the first four micromeres carry out some
RNA synthesis (Czihak, 1965; Hynes and
Gross, 1970). The experiments of Fig. 2
show clearly that these cells do not contain
typical concentrations of complex nuclear
RNAs, as do the other cells in the fourthcleavage embryo. It is important to note
that until the 16-cell stage ail the embryo
nuclei synthesize RNA at a very low rate
(perhaps 10%) relative to that observed in
the 32-cell stage and thereafter (Czihak and
Horstadius, 1970; Wilt, 1970, personal communication). Thus, the induction of the
“normal” embryonic rate of nuclear RNA
synthesis may simply be delayed by one (or
more) cell cycle in the micromeres, while it
has already occurred in at least some of the
fourth-cleavage macromeres and mesomeres.
Micromeres are essentially budded off
with a very small amount of cytoplasm from
the macromeres, and the key observation
reviewed here may be that the micromere
cytoplasm lacks the large set of nonpolysomal maternal RNAs found in other cells
of the embryo. This suggests that the micromere cytoplasm constitutes a special domain distinct from the cytoplasm of the rest
of the embryo. The micromeres may be the
first set of cells in which the embryonic
nuclei are exposed to a different environment, and this could affect the patterns of
genomic expression in the micromere nuclei
(i.e., transcription + processing). Thus the
unique fourth-cleavage division by which
the micromeres are segregated may provide
the necessary condition for the establishment of the unique pattern of function their
lineage later displays. Though the micromere cytoplasm is in some way different
from that of the rest of the embryo, its
polysomal mRNAs are likely to be the
same. As noted earlier, Tufaro and Brandhorst (1979) failed to discover any significant differences between the protein syn-
VOLUME 79,198O
thesis patterns of micromeres and those of
mesomeres + macromeres. These authors
correctly pointed out that their observations refer exclusively to the more prevalent
mRNAs. However, the same conclusion
probably pertains to the rare messages.
Whole 16-cell embryo polysomal RNA and
micromere polysomal RNA both include
about 75% of the maternal RNA sequence
set. Though not proven here, this probably
means that the same subset of rare maternal messages is being translated in 16-cell
micromeres as in mesomeres and macromeres.
Special regulators may be present in micromere cytoplasm which determine the
micromere cell lineage. We would predict
that significant differences in the micromere translational program will occur subsequently only as a result of the induction
of a unique pattern of micromere nuclear
RNA synthesis (and/or processing), rather
than as the result of a regional fourth-cleavage localization of sets of maternal message
sequences.
This research was supported by NIH Grant HD05753. The sea urchin maintenance system was partially equipped with funds supplied by NIH Biomedical Research Support Grant RR-07003, and the culture
system is maintained by NIH Grant RR-00986 from
the Division of Research Resources. S.G.E. was supported by an NIH postdoctoral fellowship.
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