HMG-D, the Drosophila melanogaster homologue of HMG 1 protein, is associated with early embryonic chromatin in the absence of histone H1.
ABSTRACT We show that HMG-D, an abundant chromosomal protein, is associated with condensed chromatin structures during the first six nuclear cleavage cycles of the developing Drosophila embryo and that histone H1 is absent from these same structures. As H1 accumulates from nuclear division 7 onwards, the nuclei become more compact and transcriptionally active. This compaction is paralleled by a reduction in size of mitotic chromatin. In addition, we find a striking correlation between the switch in HMG-D:H1 ratios and the changes that occur between nuclear cycles 8 and 13 that are collectively termed the mid-blastula transition. This transition is characterized by an increase in the nuclear cycle times, a change in the nucleo-cytoplasmic ratio, and a 5- to 20-fold decrease in nuclear volume. We propose that this is a direct consequence of a re-organization of chromatin from a less condensed state with HMG-D to a more condensed state with H1. We argue that HMG-D, either by itself or in conjunction with other chromosomal proteins, induces a condensed state of chromatin that is distinct from, and less compact than the H1-containing 30 nm fibre and that this state of chromatin could facilitate rapid nuclear cycles.
Article: A major developmental transition in early Xenopus embryos: I. characterization and timing of cellular changes at the midblastula stage.[show abstract] [hide abstract]
ABSTRACT: The Xenopus embryo undergoes 12 rapid synchronous cleavages followed by a period of slower asynchronous divisions more typical of somatic cells. This change in cell cleavage has been termed the midblastula transition (MBT). We show that at the MBT the blastomeres become motile and transcriptionally active for the first time. We have investigated the timing of the MBT and found that it does not depend on cell division, on time since fertilization or on a counting mechanism involving the sequential modification of DNA. Rather, the timing of the MBT depends on reaching a critical ratio of nucleus to cytoplasm. We view the MBT as a consequence of the titration of some substance, originally present in the egg, by the exponentially increasing nuclear material. When this substance is exhausted a new cell program is engaged, leading to the acquisition of several new cell properties.Cell 11/1982; 30(3):675-86. · 32.40 Impact Factor
The EMBO Journal vol. 13 no.8 pp. 1817 - 1822, 1994
HMG-D, the Drosophila melanogaster homologue of
HMG 1 protein, is associated with early embryonic
chromatin in the absence of histone H1
Sarbjit S.Ner and Andrew A.Travers
MRC Laboratory of Molecular Biology, Hills Road, Cambridge CB2
Communicated by A.A.Travers
We show that HMG-D, an abundant chromosomal pro-
tein, is associated with condensed chromatin structures
during the first six nuclear cleavage cycles of the
developing Drosophila embryo and that histone Hi is
absent from these same structures. As Hi accumulates
from nuclear division 7 onwards, the nuclei become more
compact and transcriptionally active. This compaction
is paralleled by a reduction in size of mitotic chromatin.
In addition, we find a striking correlation between the
switch in HMG-D:H1 ratios and the changes that occur
between nuclear cycles 8 and 13 that are collectively
termed the mid-blastula transition. This transition is
characterized by an increase in the nuclear cycle times,
a change in the nucleo-cytoplasmic ratio, and a 5- to
20-fold decrease in nuclear volume. We propose that this
is a direct consequence ofa re-organization ofchromatin
from a less condensed state with HMG-D to a more
condensed state with Hi. We argue that HMG-D, either
by itself or in conjunction with other chromosomal
proteins, induces a condensed state of chromatin that is
distinct from, and less compact than the Hl-containing
30 nm fibre and that this state of chromatin could
facilitate rapid nuclear cycles.
Key words: chromatin/embryogenesis/high mobility group 1/
histone Hl/mid-blastula transition
Chromatin remodelling during the cell cycle is crucial for
the correct regulation of genes (Grunstein, 1990; Fedor,
1992; Felsenfeld, 1992) and for the proper generation of
higher order structures (Thomas, 1984; Bradbury, 1992;
Roth and Allis, 1992). A requirement for such remodelling
is particularly apparent in the early embryo of Drosophila
melanogaster. This undergoes 13 rapid nuclear divisions in
a syncytial blastoderm, of which the first seven are
synchronous and take -8 min (Foe and Alberts, 1980).
During this phase of development the nuclei are larger and
the chromosomes under-condensed compared with nuclei of
later dividing cells. Cycles 8-13 occur at the periphery of
the embryo and have progressively longer cycle times, 9.5,
12.4 and 21.1 min respectively for divisions 11, 12 and 13
(Foe and Alberts, 1980). Correlated with the increase in the
cell cycle times there is a change in the nucleo-cytoplasmic
ratio (Edgar et al., 1986) and a 5- to 20-fold decrease in
nuclear volume between cycles 8 and 14.
The paradigm for the higher-order folding of chromatin
is the formation ofthe 30 nm fibre by the addition ofhistone
HI to core nucleosome particles (Finch and Klug, 1976;
Thoma et al., 1979; Allan et al., 1980). In Drosophila, Hi
is not detected in the early cleavage cycles (Elgin and Hood,
1973; Becker and Wu, 1992), and its first appearance has
been assumed to correspond to cycle 10 coincident with the
start of zygotic transcription (Anderson and Lengyel, 1980).
These observations imply that HI is not primarily involved
in chromatin condensation
Drosophila development. Thus, in the absence of HI and
in order for the DNA to undergo the condensation-
decondensation process during the first nine or 10 cleavage
cycles, Drosophila requires the presence ofa histone HI-like
function. However, no such protein has been identified in
Drosophila. In this paper we address the issue ofwhat other
proteins could fulfil the role ofH1 and explain the changes
that take place during early phases of development.
Circumstantial evidence suggests the vertebrate high
mobility group proteins ofthe HMG 1/2 class (Johns, 1982;
van Holde, 1989; Ner, 1992) could function in a similar
manner to Hi (Jackson et al., 1979). They have similar
properties, they stabilize and bind to bent structures (Bianchi
et al., 1992; Lilley, 1992) and have been suggested to
interact with linker DNA sequences (Johns, 1982; Schroter
and Bode, 1982). HMG-D, the Drosophila homologue of
HMG 1/2 (Wagner et al., 1992; Ner et al., 1993), is an
abundant chromatin-associated protein. It binds to DNA
cooperatively and bends DNA but exhibits little sequence
selectivity (M.E.A.Churchill and S.S.Ner, in preparation),
properties also characteristic of H1.
In our ongoing series of studies to determine the biological
role for HMG-D protein we have investigated, using an
immunological approach, whether HMG-D could,
principle, play a similar role to H1. We have determined
the spatial location ofHMG-D and HI in early embryos and
show that HMG-D is detected in all nuclei from the start
of embryogenesis, the most intense staining occurring in
mitotic chromosomes. It is also associated with other highly
condensed and transcriptionally silent chromatin structures,
such as the polar bodies and polyploid yolk nuclei. In
contrast, H1 is absent prior to nuclear cycle 7 and is not
associated with transcriptionally inert nuclei.
earliest phases of
HMG-D is present throughout embryogenesis and is
associated with all condensed chromatin structures
We have determined the temporal distribution of HMG-D
by Western blot analysis on single staged embryos. This
indicates that the protein is present at a constant level at all
stages ofdevelopment (Figure 1, data not shown for stages
9-16). Antibody staining ofembryos shows theprotein can
be detected in nuclei from the start ofembryogenesis to the
blastoderm stage. Initially the most intense staining is
© Oxford University Press
1 81 7
S.S.Ner and A.A.Travers
observed in mitotic chromosomes (Figure 3m) and the polar
bodies (Rabinowitz, 1941; Campos-Ortega and Hartenstein,
1985) (Figure 3k). Subsequently, both the polyploid yolk
nuclei and the pole cell nuclei (Figure 3j) stain strongly. The
protein persists in the latter beyond cellularization and
gastrulation. In addition, the early mitotic domains of the
3-8 . 9 10-134
Fig. 1. Western analysis of histone HI and HMG-D proteins in single
staged embryos. Total protein from single embryos (lanes 1-8, stages
1-8, respectively) were separated electrophoretically and detected after
Western transfer onto nitrocellulose with antibodies raised against
purified H1. In lanes 1-3, no protein is detected; stage 4 is the first
faint appearance of a band migrating at -33 kDa (HI has a calculated
mol. wt of 29 kDa but runs anomalously on SDS-polyacrylamide
gels). This approximately corresponds to cycles 9 and 10 when HI is
first detected in whole embryos. Stages 5-8 (lanes 5-8) show the
rapid rise in levels of HI detected. We estimate that 0.5-1 ng is
present in stage 8 embryos. Similarly, the presence of HMG-D protein
was assayed using antibodies raised against FPLC-purified protein
expressed in Ecoli. The level of protein was quantified by comparing
it with known amounts of purified protein. The amount detected
remains approximately constant (-0.2-1 ng) during stages 1-8. It
approximately doubles during stages 10-12 (data not shown). The
filter was also probed with antibodies against ca-tubulin to show that
equivalent amounts of protein had been loaded and transferred.
postblastoderm embryo are also stained by the antibody
(S.S.Ner, unpublished data).
Histone Hi is first detected in nuclear cycle 7
In contrast to HMG-D, histone HI is first detected by
Western analysis (Figure 1) on staged single embryos at
stage 4 corresponding to cleavage cycles 9 and 10. This
timing was refined by immunofluorescence-based antibody
staining which precisely localizes the first appearance ofHI
to the interphase and mitotic nuclei of some nuclear division
7 embryos where weak
(Figure 2b). In other cycle 7 embryos HI is essentially absent
(Figure 2a). Subsequently, during nuclear division cycles 8
and 9, this staining becomes more intense (Figure 2d) but,
again in contrast to HMG-D, the polyploid yolk nuclei
contain no detectable HI (Figure 3e and f). At cycle 1 the
polar bodies also lack HI (Figure 3a and b) but later become
associated with this histone (Figure 2d). After stage 7 (cycle
14), the amount of HI protein increases dramatically due
to zygotic transcription (Figure 1). We estimate stage 8
embryos contain 0.5-1
ng of HI protein (> 010
Relative levels of HMG-D and Hi change dramatically
at onset of zygotic transcription.
The average number of molecules of HMG-D per nucleus
declines with each cleavage cycle (Figure 5) although the
level ofHMG-D protein per embryo remains approximately
constant during embryogenesis. Using pure HMG-D protein
expressed in Escherichia coli as a standard, we have
calculated the relative amounts of protein present in the
Drosophila embryo. From our Western blots we estimate
that each embryo contains
(- 1-5 x 10"° molecules of HMG-D which is equivalent
in earliest embryos to
104 molecules per nucleosome
assuming -2 x 106 nucleosomes per nucleus). However,
by the cellularization stage (-5000 nuclei, cycle 14), there
in embryogenesis (>50 000 nuclei) fewer than 0.2
molecules/nucleosome. By contrast HI levels approach 1
molecule per nucleosome at approximately cycles 14 and
15 and remain at this level throughout embryogenesis.
-0.2-1 ng ofHMG-D protein
-2-5 molecules ofHMG-D per nucleosome and later
Fig. 2. Detection of histone HI before the start of zygotic transcription. HI is first detected weakly in some embryos during nuclear division cycle 7
(arrowhead in panel b). A similar cycle 7 embryo (a) shows no staining for HI. In cycle 9 (c) and early cycle 10 (d, the nuclei are in pairs and
have just completed telophase of cycle 9) the embryos begin to stain intensely for HI. Zygotic transcription starts during interphase of cycle 10
(Edgar and Schubiger, 1986). A polar body (arrow in d) which does not stain in earlier embryos (cf. Figure 3) also has histone HI associated with
Drosophila HMG-D and chromatin condensation
Thus, during the early phases of development, there is
a dramatic shift in the relative amounts ofHI and HMG-D,
with HI eventually becoming the more abundant protein
(Figure 5). This absolute increase in the level of Hi and
relative decrease in HMG-D correlates with the stages during
which nuclei start to become competent for transcription,
i.e. cycle 10 (Edgar et al., 1986; Edgar and Schubiger,
1986). Full transcriptional competence is not achieved until
cellularization, at which stage HMG-D protein is only weakly
detected. Exceptionally the pole cell nuclei stain intensely
until gastrulation (Figure 3j), correlating with the late start
of transcription in these nuclei (Edgar and Schubiger, 1986).
In addition, HMG-D, but not HI,
scriptionally silent yolk nuclei (Figure 3j).
is present in tran-
Nuclear volume and size of mitotic chromosomes
decrease after seventh nuclear division.
The nuclei in the Drosophila syncytial blastoderm change
in size during the course ofnuclear division (Foe and Alberts,
1985). We estimate, by measurement of DAPI-stained nuclei
(data not shown), that the diameter of the interphase nuclei
decreases from 4.5-5.5Amin cycles 2-8 to -2 ,im by
nuclear division 13. This 2- to 3-fold decrease in diameter
represents nearly a 20-fold decrease in nuclear volume. We
note that there is a parallel change in size of the mitotic
chromatids. We have attempted to quantify this change by
calculating the length of the chromosome on the metaphase
plate. In embryos undergoing the eighth nuclear cleavage
the chromosomes on the metaphase plate are -5Amin
length and in cycle 12 this value is 3.2 ,um (Figure 4). We
are unable to calculate how exactly this change relates to
decreases in the volume of the chromosome. However, we
feel confident that the change in nuclear volume is paralleled
by a similar reduction in the size of mitotic chromosomes
(Figure 4). We conclude that this reduction in chromosome
size reflects a greater degree of chromatin condensation.
HMG-D is a chromatin organizer
Our data show that, during the earliest phases ofDrosophila
associated with HMG-D but lack histone HI. These include
the early polar bodies, metaphase chromosomes of the
syncytial blastoderm and also the inert polyploid yolk nuclei.
With the appearance of histone HI such chromatin, as
Fig. 3. Antibody staining of staged embryo to show the presence of
HMG-D and histone HI proteins in condensed chromatin. Whole
mount immunofluorescence of nuclear division cycles 1 (a and b), 6 (c
and d) and 12 (e and f) embryos stained with DAPI (a, c and e) or
anti-HI antibody (b, d and f) to show the absence of HI in polar body
(arrow in a) and nuclei of cycle 6 embryo (d). HI begins to appear
and localize to nuclei during cycle 8 and is prominent by cycle 9 (cf.
Figure 2). At cycle 12 all nuclei stain strongly except for the yolk
nuclei (arrowhead in e). In contrast immunofluorescence staining of
embryos with anti-HMG-D antibody (h and j) shows localization of
protein in cycle 6 embryos (h), weak diffuse staining of nuclei of a
cellularizing embryo (j), weak staining of polyploid yolk nuclei
(arrowhead in j) and intense staining of the pole cells (arrow in j).
Panels g and i: the same embryos co-stained with DAPI; note how the
yolk nuclei are less intensely stained with the HMG-D antibody than
with the DAPI-stained nuclei (arrowheads in i and j), and conversely
how the pole cell nuclei are more intensely stained with the HMG-D
antibody (arrows in i and j). Panels k, I and m: higher magnification
of embryos detected for HMG-D (k and m) and histone HI using the
DAB reagent. HMG-D is present in polar body in nuclear cycle I
embryo (k) and anaphase chromosomes of cycle II (m). Panel 1: HI
in a comparable stage embryo showing essentially identical anaphase
staining pattern to HMG-D.
Fig. 4. Size of metaphase chromosomes decrease during mid-blastula
transition. Embryos were fixed and stained with antibodies against
HMG-D (brown) and a-tubulin (light brown). Embryos with the
chromatids aligned on the metaphase plate were isolated and
photographed at the same magnification. Embryos undergoing eighth
(a), 10th (b) and 12th (c) nuclear divisions. The space occupied by the
chromosome and the microtubule arrays clearly decreases with nuclear
cycle. This is illustrated in (d). Individual metaphase nuclei are
arranged adjacent to one another. The bar indicates the length of the
chromosomes on the metaphase plate. These are -5, 4.1 and 3.2Jim
respectively for the eighth, 10th and 12th nuclear cycle embryos.
S.S.Ner and A.A.Travers
Nuclear division cycle
101112 13 14A 14B 15
15 6 7 8-
Fig. 5. Summary of changes in HI and HMG-D levels during
progressive nuclear divisions and relationship to nuclear density and
volume of nuclei. The concentration of HMG-D protein remains
constant during embryogenesis but the average number of molecules
per nucleosome (1 nucleosome = 200 bp, Becker and Wu, 1992)
during nuclear divisions falls rapidly (closed circles). This value is
-1000 at cycle 6, -60 at cycle 10 and -1-5 by cycle 14. The
postblastoderm mitoses would bring this number to <0.2-0.5
molecules per nucleosome. These numbers are based on our estimates
of HMG-D proteins to be in the range 0.2-1 ng per embryo. The
data for appearance of HI are taken from our Western analysis (open
circles). Our estimates for the level of protein in stage 4 and 8
embryos indicate a rapid increase in HI, and show that the value
approximates to 1 molecule per nucleosome by late cycle 14.
Correlated with the changes in HMG-D and HI levels we depict the
changes in nucleo-cytoplasmic ratios (open squares) during division
cycles 9-13 (taken from Foe and Alberts, 1980), and the decrease in
volume of nuclei (closed squares). These values are calculated from
measuring diameters of interphase nuclei of the appropriate cleavage
cycle embryos; the volume of the nuclei decreases substantially
following migration to surface. The diameter of nuclei decreases by
half during cycles 8-13. The correlation of these changes with zygotic
transcription is also indicated. Full zygotic transcription is attained
during cycle 14.
exemplified by the size of the mitotic chromatids, becomes
progressively more compact until eventually HMG-D is
supplanted by the histone. We conclude that the organization
of early embryonic chromatin differs from that characteristic
of later developmental stages. Other evidence for the
existence of a different state of early embryonic chromatin
has been presented by Becker and Wu (1992) who observed
that in chromatin reconstituted from early embryonic extracts
the nucleosome spacing is
of HI this value increases to -200 bp.
In the early embryo HMG-D is highly abundant. It is
deposited in the egg by the mother but thereafter is
maintained at an approximately constant level per embryo.
Consequently, with each nuclear division the average number
ofHMG-D molecules per nucleus falls while during nuclear
cycles 7-14 the amount of HI rapidly increases. These
observations are compatible with HMG-D, either by itself
or in conjunction with other chromosomal components,
providing a role in the condensation of nucleosomes into
-180 bp whereas on addition
higher order structures in the early embryo and thus may
perform a function analogous to that of histone H1.
A structural role for HMG-D?
The abundance ofHMG-D and its properties are consistent
with a structural role for this protein. The higher-order
structures produced in the presence of HMG-D would be
distinct from the chromatin containing HI. Such a role for
HMG-D is also consistent with the observed properties of
the vertebrate homologues HMG
proteins bend DNA (Paull et al., 1993; Pil et al., 1993),
can interact directly with HI in vitro (Carballo et al., 1983;
Kohlstaedt et al., 1987) and, under physiological conditions,
facilitate the in vitro formation of complex nucleoprotein
structures, including both nucleosomes (Bonne-Andrea
et al., 1984) and invertasomes functional for site-specific
recombination (Paull et al., 1993).
Histone HI is required both for the formation of the 30
nm fibre and for the further condensation of chromatin into
more compacted structures (Allan et al., 1980; Hill et al.,
1991). The former involves both the sealing ofthe two gyres
of nucleosomal DNA by the bivalent globular domain ofHI
and also cooperative interactions between adjacent HI
molecules (Clark and Thomas, 1988), while the N-terminal
domain is necessary for the latter process (Hill et al., 1991).
Although the precise disposition of HI in the 30 nm fibre
is not established
properties ofHMG-D, that the latter protein could fulfil one
or more functions of HI. In particular, HMG-D possesses
a region rich in alanine and lysine residues which is similar
to the C-terminal domain ofHI (Ner et al., 1993). In certain
other organisms (sea urchin and Xenopus) the early embryo
contains a homologue ofHI termed 'cleavage cycle' histone
HI (Levy et al., 1982; Smith et al., 1988; van Holde, 1989)
that replaces the normal somatic H1. In Xenopus this is
termed B4 and is restricted to early development (Dimitrov
et al., 1993). In these HI variants homology to the somatic
HI is largely limited to the globular domain, with both the
N- and C-terminal domains being deficient in basic residues.
Although no such variants have so far been identified in
Drosophila it remains possible that such a protein may also
be necessary for the condensation of early embryonic
1 and HMG 2. These
it seems plausible, from the known
What could be the physiological significance of different
forms of condensed chromatin? In principle the structures
containing HMG-D and HI could represent alternative
modes ofcompaction. A second possibility is that the HMG-
D-containing structure is a less compacted intermediate on
the pathway to solenoid formation. In either view, a looser
structure could form in the absence ofHI and could facilitate
the rapid condensation and decondensation required during
the very short early cleavage cycles.
As a consequence of the different condensed chromatin
structures generated and changes in HMG-D and HI levels
a model can be proposed which also explains the observed
changes associated with mid-blastula transition (MBT,
Newport and Kirschner, 1982). The appearance of HI is
correlated with a decrease in the size of nuclei and the
metaphase chromosome as well as a lengthening of nuclear
division times, and the acquisition of transcriptional
competence during cycle 10 (Anderson and Lengyel, 1980;
Drosophila HMG-D and chromatin condensation
Edgar and Schubiger, 1986). In general, nuclei which stain
strongly for HMG-D are transcriptionally inactive. For
example, yolk nuclei remain transcriptionally inactive and
gastrulation has begun. These observations strongly suggest
that the chromatin generated in the presence of HMG-D is
transcriptionally silent, and lead us to argue that transcription
only begins when HI levels have reached a particular
threshold value and overcome the HMG-D effects, such as
interphase or the nucleo-cytoplasmic ratio (Edgar et al.,
1986). Nuclei of cycle 8 and 9 embryos are unable to
transcribe genes even when interphase is extended through
the use of cell cycle inhibitors (Edgar and Schubiger, 1986).
Therefore, we suggest that the ratio of H1/HMG-D and the
extent of remodelled chromatin are the crucial determinants
in the acquisition of transcriptional competence. Similar
observations have been described in the Xenopus system in
which B4, the H1 variant, disappears during MBT and there
is a correlated change in the accessibility of embryonic
chromatin to class III transcriptional machinery (Dimitrov
et al., 1993).
The first appearance of HI during cycles 7 and 8 is
significantly earlier than previously estimated (Becker and
Wu, 1992) and implies that HI must arise from translation
of the maternally deposited mRNA (Ruddell and Jacobs-
Lorena, 1985) since zygotic transcription of the HI gene
begins during cycle 10 (Anderson and Lengyel, 1980). If
this implication is correct it follows that translation of this
RNA is tightly regulated to start at cycles 7 and 8. It is during
cycle 7 that the size of the nuclei begins to decrease. By
cycles 10-12 a sufficient amount of histone HI has
accumulated to allow the reorganization of chromatin to a
transcriptionally active state. Subsequently, increased zygotic
transcription elevates histone H 1
exponential increase together with the increased number of
nuclei rapidly deplete HMG-D protein to levels that cannot
have global effects on chromatin structure. Thus,
structures where HI is absent and HMG-D is present, such
as yolk nuclei, or in structures where HMG-D protein
persists for longer periods, such as in pole cells, we suggest
HMG-D suppresses transcription. We note that HMG-D is
present in later stages of embryogenesis as is HMG-Z, a
related protein which has extensive sequence homology to
HMG-D (Ner et al., 1993). The combined levels of these
proteins during the later stages of embryonic and larval
development remain relatively constant and do not exceed
the values per nucleosome for HI (S.S.Ner, unpublished).
Hence, we suggest HMG-D and HMG-Z have a second
function analogous, perhaps, to that of HI in organizing
chromatin locally into active or inactive domains.
Our future efforts are directed towards binding studies of
HMG-D to nucleosomes and competition experiment with
HI to provide biochemical evidence in support of the model
proposed in this study.
is not purely a function of the length of
Materials and methods
Generation of antibodies against HMG-D
Full-length HMG-D was overproduced in E.coli and purifiedto homogeneity
(M.A.Searles and S.S.Ner, unpublished). Polyclonal antibodies were raised
in rabbits against this protein. IgG-containing fractions were isolated by
affinity purification from a proteinA-Sepharose column (HarlowandLane,
1989). These fractions were preadsorbed against a large volume of0-20 h
old fixed Drosophila embryos prior to use in Western analysis and for embryo
Single embryo Western analysis
Embryos were staged under a dissecting microscope (Campos-Ortega and
Hartenstein, 1985) and placed on the inverted lid of an Eppendorftube which
contained 2 1l of standard SDS gel loading buffer. The embryowas crushed
by placing upon it a small cut piece of coverslip and pressing gently with
a pair of forceps. The lid was then replaced on the tube and spun in a
centrifuge for 30 s. A further 5 1l of SDS loading buffer was added, and
the sample was boiled and loaded onto a 15% SDS-polyacrylamidemini-
gel. Electrophoresis and Western transfer were performed using standard
conditions. The protein bands were detected using secondary antibodies
coupled with horseradish peroxidase and a chemiluminescence detection
Procedures for fixation and antibody staining of embryos were carried out
as described (Lawrence and Johnston,
antibodies (Amersham) were either conjugated with biotin and the antigens
visualized with the Vectastain Elite ABC system (Vector Laboratories), or
conjugated with fluorescein isothiocyanate (FITC) and the antigens detected
using epifluorescence optics. The embryos that were stained with the FITC-
conjugated secondary antibody were counterstained with the nuclear dye
1989). Anti-rabbit secondary
We thank Rob Kay, Hugh Pelham, Mair Churchill and John Girdlestone
for helpful comments on the manuscript; Andrew Belmont (University of
Illinois) for pointing out the absence ofHI in early embryos; Teresa Langford
for immunization and welfare of animals; J.Kadonaga and R.Kamakaka for
providing a generous sample of anti-HI serum; and the members of the
photography department in the Laboratory of Molecular Biology for help
with figures and photographs.
Allan,J., Hartman,P.G., Crane-Robinson,C. and Aviles,F.X. (1980) Nature,
Anderson,K.V. and Lengyel,J.A. (1980) Cell, 21, 717-727.
Becker,P.B. and Wu,C. (1992) Mol. Cell. Biol., 12, 2241-2249.
Bianchi,M.E., Falciola,L., Ferrari,S. and Lilley,D.M.J. (1992) EMBO J.,
Bonne-Andrea,C., Harper,F., Sobczak,J. and De Recondo,A.-M. (1984)
EMBO J., 3, 1193-1199.
Bradbury,E.M. (1992) Bioessays, 14, 9-16.
Campos-Ortega,J.A. and Hartenstein,V. (1985) The Embryonic Development
of Drosophila melanogaster. Springer-Verlag, Berlin.
Clark,C.D. and Thomas,J.O. (1988) Eur. J. Biochem., 178, 225-233.
Dimitrov,S., Almouzni,G., Dasso,M. and Wolffe,A.P. (1993) Dev. Biol.,
Edgar,B.A. and Schubiger,G. (1986) Cell, 44, 871-877.
Edgar,B.A., Kiehle,C.P. and Schubiger,G. (1986) Cell, 44, 365-372.
Elgin,S.C.R. and Hood,L.E. (1973) Biochemistry, 12, 4984-4991.
Fedor,M.J. (1992) Curr. Opin. Cell Biol., 4, 436-443.
Felsenfeld,G. (1992) Nature, 355, 219-223.
Finch,J.T. and Klug,A. (1976) Proc. NatlAcad. Sci. USA, 73, 1897-1901.
Foe,V.E. and Alberts,B.M. (1980) J. Cell Sci., 61, 31-70.
Foe,V.E. and Alberts,B.M. (1985) J. Cell Biol., 100, 1623-1636.
Grunstein,M. (1990) Annu. Rev. Cell. Biol., 6, 643-678
Harlow,E. and Lane,D. (1988) Antibodies. A LaboratoryManual. Cold
Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
Hill,C.S., Rimmer,J.M., Green,B.N., Finch,J.T. andThomas,J.O. (1991)
EMBO J., 10, 1939-1948.
Johns,E.W. (ed.) (1982) The HMG Chromosomal Proteins. Academic Press,
Kohlstaedt,L.A., Sung,E.C., Fujishige,A.and Cole,R.D. (1987)J. Biol.
Chem., 262, 524-526.
Lawrence,P.A. and Johnston,P. (1989) Development, 105, 761-767.
Levy,S.,Sures,I. andKedes,L. (1982)J. Biol. Chem., 257, 9438-9443.
(1983) EMBO J.,