Download The nullo protein is a component of the actin

1863
Journal of Cell Science 107, 1863-1873 (1994)
Printed in Great Britain © The Company of Biologists Limited 1994
The nullo protein is a component of the actin-myosin network that mediates
cellularization in Drosophila melanogaster embryos
Marya A. Postner and Eric F. Wieschaus*
Department of Molecular Biology, Princeton University, Princeton, NJ 08544, USA
*Author for correspondence
SUMMARY
After the 13th nuclear division cycle of Drosophila embryogenesis, cortical microfilaments are reorganized into a
hexagonal network that drives the subsequent cellularization of the syncytial embryo. Zygotic transcription of the
nullo and serendipity-α genes is required for normal structuring of the microfilament network. When either gene is
deleted, the network assumes an irregular configuration
leading to the formation of multinuceate cells. To investigate the role of these genes during cellularization, we have
made monoclonal antibodies to both proteins. The nullo
protein is present from cycle 13 through the end of cellularization. During cycle 13, it localizes between interphase
actin caps and within metaphase furrows. In cellularizing
embryos, nullo co-localizes with the actin-myosin network
and invaginates along with the leading edge of the plasma
membrane. The serendipity-α (sry-α) protein co-localizes
with nullo protein to the hexagonal network but, unlike the
nullo protein, it localizes to the sides rather than the
vertices of each hexagon. Mutant embryos demonstrate
that neither protein translationally regulates the other, but
the localization of the sry-α protein to the hexagonal
network is dependent upon nullo.
INTRODUCTION
taposition: the sides of each polygon in the array are formed
by converting the ‘fuzzy’ actin organization at the cap margins
into more finely aligned actin filaments (Simpson and
Wieschaus, 1990). Each hexagonal interface of the actin
network defines the site of membrane invagination. Cytoplasmic myosin co-localizes with actin in the hexagonal network
(Warn et al., 1980; Young et al., 1991), and both filamentous
actin and functional cytoplasmic myosin are required for
membrane invagination (Zalokar and Erk, 1976; Foe and
Alberts, 1983; Kiehart et al., 1990). Contraction of this
actin/myosin array has been postulated to provide a mechanistic force driving the invagination. This role for the actinmyosin network is based in part on an analogy with the ‘contractile rings’ of actin and myosin that are thought to drive
invagination of the plasma membrane during conventional
cytokinesis (Mabuchi, 1986; Salmon, 1989; Schroeder, 1990;
Satterwhite and Pollard, 1992).
Most of the components of the hexagonal array are supplied
by maternal transcription during oogenesis and are already
present as RNA or protein in the unfertilized egg. Cellularization, however, marks the point in Drosophila development
when the embryo becomes dependent on gene products
supplied by the embryo’s own transcription (Arking and
Parente, 1980; Edgar and Schubiger, 1986). A small number of
genes have been identified whose zygotic products are required
for the formation of a normal actin array (Wieschaus and
Sweeton, 1988; Merrill et al., 1988). Embryos lacking either the
nullo or the serendipity-alpha (sry-α) gene show very similar
In Drosophila embryos, the early nuclear divisions are not
followed by cytokinesis and the embryo initially develops as a
syncytium. This organization persists until after the 13th
division, at which time the embryo consists of approximately
6,000 nuclei arranged in a monolayer in the embryo’s cortex.
Subdivision of the cortical cytoplasm into individual cells is
known as cellularization. During this process, plasma
membrane invaginates from the surface in a roughly hexagonal
pattern, precisely separating each nucleus from its immediate
neighbors. Once the membrane has reached a depth of about
25 µm, the base of the invaginating membrane furrow begins
to widen, eventually separating the newly formed cells from
the underlying yolk. The resulting cellular blastoderm consists
of a single layer of columnar cells surrounding the central yolk
sac.
During the first ten minutes of cycle 14, a highly organized
array of F-actin is formed on the cytoplasmic face of the
invaginating plasma membrane (Fig. 1, see also Warn and
Magrath, 1983; Simpson and Wieschaus, 1990; Warn and
Robert-Nicoud, 1990; Schejter and Wieschaus, 1993). Prior to
formation of the array, the cortical actin of the embryo is
organized in ‘caps’ overlying each nucleus. Initially the caps
formed in cycle 14 resemble those seen in the preceding interphases. However, in contrast to earlier caps, which remain
static during interphase, the cycle 14 caps soon enlarge until
their bases touch. The actin array arises in regions of cap jux-
Key words: Drosophila embryo, cytokinesis, contractile ring
1864 M. A. Postner and E. F. Wieschaus
abnormalities in the actin array: some of the sides of the
hexagons are unusually thick while others are extremely thin or
missing altogether (Fig. 1B,C). In nullo embryos, the initial few
minutes of network formation appear normal (Wieschaus and
Sweeton, 1988; Simpson and Wieschaus, 1990). However, at
the onset of membrane invagination, network formation is
incomplete and the uneven distributions and sporadic disruptions in the actin-myosin network become obvious. Once
underway, network and membrane invagination appear to
proceed normally: neither the kinetics of membrane extension
nor the length of the newly formed cells is significantly different
from that observed in wild-type embryos. However, cleavage
furrows do not invaginate where the network is discontinuous
and multinucleate cells form. The only obvious difference
between nullo and sry-α mutant embryos is that sry-α embryos
have fewer multinucleate cells (Merrill et al., 1988;
Schweisguth et al., 1990; E. Schejter, personal communication).
Molecular characterization of the nullo and sry-α genes has
revealed that both genes encode single, blastoderm-specific
transcripts that are uniformly distributed throughout the
syncytial embryo and accumulate in large amounts over a short
period of time (Vincent et al., 1985; James and Vincent, 1986;
Rose and Wieschaus, 1992). Transcript levels reach a sharp
peak around the onset of cellularization and subsequently
decrease in a rapid, spatially patterned manner. The main difference in the transcription pattern of the two genes is that sryα transcripts arise, peak and decline slightly later than nullo.
Neither gene is required for the transcription of the other (Rose
and Wieschaus, 1992).
The sry-α protein is 58 kDa in size, lacks extensive
homology to any known proteins and shows few structural
motifs (Ibnsouda et al., 1993). Immunolocalization indicated
that during cellularization sry-α protein localizes to the leading
edge of the invaginating plasma membrane (Schweisguth et al.,
1990). Like its transcript, the sry-α protein is short-lived. The
nullo gene is predicted to encode a 23 kDa protein lacking
homology to known proteins, including the sry-α protein.
Sequence analysis demonstrated that the nullo protein has an
excess of basic amino acids (predicted pI is 11.4) and
suggested that the protein may be myristoylated (Rose and
Wieschaus, 1992). However, previous studies did not address
intracellular localization of the nullo protein or its specific cell
biological function during cellularization.
While many broad questions regarding the mechanisms of
cellularization remain unanswered, several specific questions
about the nullo and sry-α proteins are amenable to experimentation: do the proteins directly interact with the cellularization machinery or are they indirect participants in cellularization? What are the functions of the nullo and sry-α proteins
during the process? Is the similarity of their mutant phenotypes
due to their participation in a common pathway or do the two
proteins function independently? To address these questions,
we have generated monoclonal antibodies to both nullo and
sry-α proteins, and examined their distribution during cellularization in both wild-type and mutant embryos.
MATERIALS AND METHODS
Genotypes and stocks used
Ore-R was used as the wild-type stock. Embryos with the nullo
phenotype were collected from balanced stocks containing deficien-
cies that uncover the nullo locus. The deficiencies Df(1)6F1-2 and
Df(1)LIMDF were most commonly used (for description see Simpson
and Wieschaus, 1990; Rose and Wieschaus, 1992). Embryos with the
sry-α phenotype were collected from a stock that is heterozygous for
Df(3R)X3F, which uncovers the sry-α gene. Since no point mutations
exist for either gene, the mutant phenotype only arises in deficiency
embryos. The terms ‘nullo mutant’ and ‘sry-α mutant’ are used to
describe the deficiency embryos.
Production and screening of monoclonal antibodies
Monoclonal antibodies to the nullo protein were generated using a
nullo-glutathione S-transferase fusion protein as the antigen. An inframe fusion of the entire nullo protein to the carboxyl terminus of
glutathione S-transferase (Smith and Johnson, 1988) was constructed
in the following manner. The nullo coding region was PCR amplified
from the nullo M1 cDNA (Rose and Wieschaus, 1992) using primers
homologous to the ends of the coding region. The primers also
contained an external stretch of bases that lacked homology to nullo
and contained an EcoRI restriction site. The resulting PCR product
was digested with EcoRI and ligated into the EcoRI site of pGEX-2T
(Pharmacia). The orientation of the inserts was determined by restriction mapping.
The pGEX-2T-nullo plasmid was transformed into Escherichia coli
strain JM101 and the production of fusion protein was induced with
1 mM IPTG (Pharmacia). After 3 hours, the cells were harvested.
Because the fusion protein was stubbornly insoluble, it was excised
from a preparative acrylamide gel, electroeluted with Elutrap
(Schleicher and Schuell), and dialyzed against MTPBS (Smith and
Johnson, 1988). The purified protein was used to immunize two mice
and monoclonal antibodies were produced following standard
protocols (Harlow and Lane, 1988). Supernatants from the monoclonal
lines were tested by western blot for recognition of the fusion protein
and of glutathione S-transferase. The 23 lines that recognized only the
fusion protein were tested by western blot for reactivity with proteins
from two- to three-hour Ore-R embryos. Monoclonal supernatants
5C3-12 and 2F8-18 specifically recognize the nullo proteins. Because
the 5C3-12 antibody reacts more strongly with the nullo proteins than
does 2F8-18, it was used preferentially unless otherwise indicated.
Monoclonal antibodies to the sry-α protein were generated using a
truncated version of the sry-α protein as the antigen. This protein,
which contains amino acids 46 to 530 of sry-α, was produced from a
T7 RNA polymerase-inducible vector (Studier and Moffat, 1986). The
plasmid, pPαNN, contains a NarI to NcoI fragment of the sry-α gene
cloned in pET3a. It was generously provided by Alain Vincent. The
plasmid was transformed into E. coli strain BL21-Lys S. Fusion
protein production was induced for three hours with 1 mM IPTG. The
cells were lysed by freezing with 3.3 mg/ml lysozyme and then
thawing. Inclusion bodies were purified by isolating all proteins
insoluble in DOC buffer (200 mM NaCl, 1% sodium deoxycholate,
1% NP40, and 1 mM DTT). The protein pellet was washed three times
with 0.5% Triton X-100, 1 mM EDTA and 1 mM DTT, before being
resuspended in TE (10 mM Tris, 1 mM EDTA). This protein preparation was greatly enriched for the truncated sry-α protein. Two mice
were immunized with it and monoclonal antibodies were produced.
The monoclonal supernatants were screened by western blot to test
reactivity with the truncated sry-α protein that served as the antigen.
Reactive supernatants were further tested for reactivity with a 58 kDa
protein from two- to three-hour wild-type embryos. Monoclonal
supernatants 1G10, 3H6, 4G4, 6B12 and 6F4 all react strongly with
the sry-α protein. Antibody 1G10 was used in most instances.
Immunoprecipitation
Two- to three-hour wild-type embryos were dechorionated, ground in
80 mM Tris with 2% SDS, and boiled for five minutes. The extracts
were chilled on ice. Modified RIPA buffer (50 mM Tris, 300 mM
NaCl, 1% NP40, 0.5% sodium deoxycholate) and 10% Triton X-100
were added to the extracts to achieve a final concentration of 0.2%
nullo protein in Drosophila embryos 1865
SDS and 1% Triton X-100. Samples of 75 µl each of nullo monoclonal antibodies 5C3-12 and 2F8-18 were incubated with the protein
extracts for one hour at 4˚C. Then, 50 µl of 50% Protein A-Sepharose
beads (Pharmacia) in RIPA buffer (above recipe plus 1% SDS) were
added and incubated at 4˚C. After one hour, the pellet was recovered
and washed twice in RIPA buffer.
Antibody staining of embryos
Two- to three-hour embryos were dechorionated and fixed by one of
two procedures: (1) fixation for 20 minutes with 18.5% formaldehyde
saturated with heptane followed by manual devitellinization or (2)
boiling for 10 seconds in Triton-salt solution (68.4 mM NaCl, 0.03%
Triton X-100) followed by the addition of a vast excess of ice-cold
Triton-salt solution, devitellinization using methanol and heptane, and
post-fixation of at least one hour in methanol. The fixed and
devitellinized embryos were incubated for one hour at room temperature with PBT10 (PBS with 10% BSA and 0.1% Tween-20). Incubations with the primary antibodies were performed overnight at 4˚C:
sry-α monoclonal antibody 1G10 was diluted 1:25 in PBT1 (PBS with
1% BSA and 0.1% Tween-20); nullo monoclonal 5C3-12 was diluted
1:15 in PBT1; antisera to cytoplasmic myosin (kindly provided by
Dan Kiehart) was diluted 1:250 in PBT1. After incubation with the
appropriate primary antibody, the embryos were washed once with
PBT1 and four times for 30 minutes each with BNT100 (PBS with
2% normal goat serum, 100 mM NaCl, 1% BSA, and 0.1% Tween20). Preabsorbed fluorescent secondary antibodies were diluted 1:250
in PBT0.1 (PBS with 0.1% BSA and 0.1% Tween-20) and embryos
were incubated with them at room temperature for one to three hours.
To visualize filamentous actin, embryos were stained for 20 minutes
with either 0.165 µM bodipy-phalloidin or 0.165 µM rhodamine-phalloidin (Molecular Probes). After several washes in PBS-Triton, they
were incubated for 3 minutes with 1 µg/ml Hoechst 33258 (Polysciences), a DNA-specific dye. The embryos were washed extensively
in PBS-Triton and PBS before being mounted in Aquapolymount
(Polysciences). Embryos were examined and photomicrographs made
using a Bio-Rad MRC600 confocal microscope.
Fig. 1. The actin-myosin network that forms in Drosophila embryos
during cycle 14. In wild-type embryos (A), the array consists of
approximately 6000 polygons of roughly equal size. Each polygon
defines the area above a single somatic nucleus. Bar, 100 µm. At
higher magnification, the wild-type array (B) shows a very regular
configuration of polygons composed of uniformly thick actin
interfaces. In embryos deficient for nullo (C) or sry-α (D), the array
has interfaces of irregular thickness and the individual polygons are
of variable size. Actin microfilaments were visualized by staining
with FITC-labeled phallacidin. Bar in D (also applies to B and C),
10 µm.
RESULTS
nullo protein is blastoderm-specific
In order to characterize the distribution of nullo protein, monoclonal antibodies to the protein were isolated. The antigen
was a fusion protein consisting of glutathione S-transferase and
the full-length nullo protein. Supernatants from two monoclonal lines recognized proteins from two- to three-hour-old
embryos in the size range predicted for nullo (approximately
23 kDa). Both supernatants reacted with the same protein
doublet of 25 and 26.5 kDa (Fig. 2A). The doublet was absent
in protein preparations from embryos homozygous for a small
deficiency of nullo (Fig. 2B and C), indicating that both
proteins are encoded by the nullo gene and that both monoclonal lines are specific for nullo proteins. The difference in
migration between the two forms of nullo is probably due to a
post translational modification, since northern blots and
sequence analysis predict only a single nullo product (Rose and
Wieschaus, 1992).
The temporal profile of nullo protein levels was determined
by western blot analysis of single Ore-R embryos staged prior
to homogenization (Fig. 1D). nullo protein was detectable
between interphase of cycle 13 and the beginning of gastrulation. The amount of nullo protein is low in cycle 13 interphase
embryos, increases greatly during the 13th division, and
reaches an apparent peak in early cycle 14. Levels of nullo
protein remain high during the initial slow phase of cellularization, when invagination of membrane is thought to depend
on the incorporation of new membrane behind the furrow
(Turner and Mahowald, 1976). Levels drop rapidly during the
subsequent ‘fast’ phase and protein is barely detectable by the
beginning of gastrulation. Both forms of nullo protein show
similar kinetics of accumulation and disappearance. The
pattern parallels that previously reported for the nullo RNA
with a lag of about fifteen to twenty minutes.
Intracellular localization of nullo proteins
Antibodies to nullo were used for immunolocalization of the
protein during syncytial and cellular blastoderm stages. To
visualize the microfilament network independently, wild-type
embryos were simultaneously stained with antibodies against
myosin or with phalloidin. nullo protein is first detectable in
whole-mount embryos during interphase of cycle 13 (Fig. 3A).
At this stage, all detectable nullo protein is localized in the
cortical cytoplasm of the embryo and is punctate or vesicular
in nature. The protein is restricted to a region apical to the
monolayer of nuclei, and appears to be associated with the
plasma membrane. It does not co-localize with the actin caps
that form above the interphase nuclei and instead appears to be
restricted to the areas between the caps. When viewed from the
surface, the resulting pattern of nullo distribution resembles a
1866 M. A. Postner and E. F. Wieschaus
Fig. 2. Western blots of embryo extracts using
nullo monoclonal antibody. (A) Extracts from
2- to 3-hour wild-type embryos. Lane 1
contains approximately 10 embryos; lane 2
contains approximately 5 embryos; and lane M
contains unstained low molecular mass
markers (Bio-Rad). The monoclonal antibody
recognizes two proteins of approximately 25
and 26.5 kDa. (B) Each lane contains proteins
extracted from a single cycle 14 embryo. The
embryos were collected from a cross in which
one-quarter of the embryos are deleted for the
nullo locus. The embryos were randomly
harvested during early cycle 14 and no attempt
was made to pick normal or mutant embryos.
The embryos in lanes 2, 4, 8 and 11 lack both
nullo proteins and are assumed to be deleted
for the nullo locus. (C) In this control
experiment, each lane contains proteins
extracted from a single wild-type embryo in
cycle 14. As expected, two forms of the nullo
protein were recovered from all embryos. (D)
Wild-type embryos were carefully staged and
harvested at precise stages, from the beginning
of cycle 13 through early gastrulation. Each
lane contains protein from a single embryo. Lanes 1 and 2, cycle 13, interphase; lanes 3 and 4, cycle 13, mitosis; lanes 5 and 6, cycle 14, precellularization; lanes 7 and 8, slow phase of cellularization; lane 9, beginning of fast phase of cellularization; lanes 10 and 11, end of fast phase;
and lanes 12 and 13, gastrulation. The levels of the nullo protein are highest from the 13th division through the slow phase of cellularization.
relatively regular network of interlocking rings that encompasses the entire embryo (Fig. 3A). The intensity of staining
with the nullo antibody is weak at this stage.
During the mitosis of cycle 13, nullo protein is localized
within the pseudocleavage furrows that transiently invaginate
between the dividing nuclei. Although actin underlies the
entire plasma membrane during this stage, nullo appears to be
associated only with invaginated membrane regions. When
viewed from above (Fig. 3B), nullo forms elongated hexagons,
each enclosing a mitotic nucleus. The intensity of nullo
antibody staining at this stage is considerably greater than that
in interphase of cycle 13.
At the onset of cycle 14, when actin caps re-form and the
hexagonal array of actin and myosin begins to resolve, nullo
protein localizes to the bases of the caps (Fig. 4A,B,C,D).
During the next ten minutes, as the network matures, nullo
protein is colinear with the network and shows an intense and
orderly punctate staining pattern (Fig. 4E,F,G,H). Closer
analysis of the staining patterns within each hexagonal unit
reveals that nullo and actin are distinguishable. When visualized with phalloidin or phallacidin, the hexagonal units are
defined by uninterrupted ‘lines’ of filamentous actin. The ends
of these lines meet to form the hexagon’s vertices (Rose and
Wieschaus; see also Fig. 5). By comparison, nullo protein
shows a discontinuous, punctate staining pattern. Many dots of
nullo protein are aligned to give the hexagonal pattern. While
there is nullo present along the sides of each hexagon, the
highest concentration of nullo staining occurs at the vertices.
In addition, some nullo protein is present in the apical regions
above the nuclei, as well as in the cytoplasm just below the
Fig. 3. Localization of the nullo protein in late syncytial embryos. When wild-type embryos at interphase of cycle 13 (A) are viewed from
surface, the nullo protein forms a hexagonal network. The nullo protein localizes to the pseudocleavage furrows during the mitosis of cycle 13
(B). It forms a hexagonal array in cycle 14 (C). The absolute level of staining with the nullo antibody dramatically increases from interphase of
cycle 13 to early cycle 14. To compare spatial patterns of distribution, all panels have been printed at the same intensity levels. Bar in C (also
applies to A and B), 10 µm.
nullo protein in Drosophila embryos 1867
Fig. 4. Cellularizing wild-type embryos double labelled with the nullo antibody (A,C,E,G,I,K,M,O,Q,S) and cytoplasmic myosin antibody
(B,D,F,H,J,L,N,P,R,T). The first and second columns show surface views, the third and fourth columns show cross-sections from the same
embryos. During conversion of the actin caps to a hexagonal network (A-D), the nullo protein is localized to the forming network (A,C). It
maintains a localization to the leading edge of the invaginating membrane at the initiation of membrane invagination (E-H) and during slow
phase (I-L). During the fast phase of cellularization (M-P), the levels of nullo protein present at the cellularization front decrease and the
protein becomes dispersed throughout the cytoplasm. By the completion of cellularization (Q-T) nullo protein is almost undetectable. Bar in D
(applies to entire figure), 10 µm.
1868 M. A. Postner and E. F. Wieschaus
bases of the nuclei, where its staining pattern gives the impression of being vesicular.
During the slow phase of cellularization (Fig. 4I,J,K,L),
nullo protein continues to be primarily associated with the
plasma membrane at and slightly apical to the invaginating
furrow canal, where it maintains its initial distribution in a
hexagonal array. In some confocal cross-sections, nullo
staining appears as a line extending apically from the furrow
canal. This line never extends more than half-way to the apical
plasma membrane. Unlike actin, nullo is not localized to the
non-invaginated plasma membrane on the apical surface,
although some slight staining above background is observed in
Fig. 5. Cellularizing wild-type embryos double labelled with the sry-α antibody (A,C,E,G,I,K,M,O) and phalloidin (B,D,F,H,J,L,N,P). The first
and second columns show surface views; the third and fourth columns show cross-sections from the same embryos. During actin-myosin
network resolution (A-D) and during slow phase of cellularization (E-H), sry-α protein localizes to the hexagonal array (see arrows in C and
G). The levels of localized sry-α protein remain high at the end of slow phase (I-L) but decrease during the latter half of fast phase (M-P). The
protein becomes more dispersed in the cytoplasm (M, O), although some staining remains in a hexagonal pattern (M). sry-a protein also
localizes in a sphere above each nucleus (G and K, arrowheads) until the final stages of cellularization (G,K,) as well as maintaining a more
general localization in the apical cytoplasm (G,K). Bar in D (applies to entire figure), 10 µm.
nullo protein in Drosophila embryos 1869
the cytoplasm apical to the nuclei. All cytoplasmic staining
observed at these stages is still punctate in appearance.
As cellularization progresses into the fast phase (Fig.
4M,N,O,P), nullo protein maintains its localization to the cellularization front, but the intensity of staining is significantly
diminished and more variable from embryo to embryo than in
previous stages of cellularization. When viewed tangentially at
the level of the furrow canals, the nullo protein continues to
form a thin network of interconnecting hexagons. This is in
contrast to the actin-myosin network, which thickens, resulting
in a ring-like appearance of the individual units in the network.
During the course of fast phase (Fig. 4Q,R,S,T), progressively
Fig. 6. Localization of the nullo proteins in nullo and sry-α mutant embryos. Embryos deleted for nullo locus (E,F,G,H) and their heterozygous
siblings (A,B,C,D) were double labelled using nullo antibody (A,C,E,G) and cytoplasmic myosin antisera (B,D,F,H). The specificity of the
nullo antibody is demonstrated by its failure to stain nullo mutant embryos (E,G). In embryos lacking sry-α, (M,O), the nullo protein forms a
network that is collinear with the actin-myosin hexagonal array: a relatively normal nullo network is seen everywhere except where the actinmyosin network is disrupted. Bar in D (applies to entire figure), 10 µm.
1870 M. A. Postner and E. F. Wieschaus
less nullo protein is localized to the furrow canal and the
majority of nullo protein becomes dispersed throughout the
cytoplasm. The intensity of nullo staining continues to decline
and is no longer detectable above background when cellularization is completed.
Intracellular localization of the serendipity-alpha
protein
Since nullo and serendipity-alpha (sry-α) mutants have a
common phenotype and both proteins have now been reported
to localize to the hexagonal array of actin, a detailed compar-
Fig. 7. Localization of the sry-α protein in nullo and sry-α mutant embryos. Comparison of embryos deleted for sry-α (E,F,G,H) and their
heterozygous siblings (A,B,C,D) when double labelled with sry-α antibody (A,C,E,G) and phalloidin (B,D,F,H) confirm that the sry-α antibody
is specific: sry-α heterozygotes have a hexagonal array of sry-α protein (A,C) whereas their mutant siblings show no such staining (E,G).
Embryos deleted for nullo (M,N,O,P) and their heterozygous siblings (I,J,K,L) were also double-labelled with sry-α antibody (I,K,M,O) and
phalloidin (J,L,N,P). The sry-α protein in nullo mutants (M,O) does not form a hexagonal network at the level of the cellularization front. Bar
in D (applies to entire figure), 10 µm.
nullo protein in Drosophila embryos 1871
ison of the intracellular distributions of these two proteins is
merited. Mice were immunized with a truncated version of the
sry-α protein and supernatants from several of the resulting
monoclonal lines reacted with a protein from two- to threehour fly embryos of approximately 58 kDa, the size of sry-α
as defined by polyclonal antisera (Schweisguth et al., 1990).
The specificity of the monoclonal antibodies was shown by the
failure of embryos deleted for the sry-α gene to exhibit any of
the staining patterns described (see Fig. 7E,G).
Although the sry-α protein has been reported to be present
in cycle 13 embryos (Schweisguth et al., 1990), our antibodies
only detect sry-α protein above background staining during the
formation of the hexagonal network in early cycle 14. As soon
as the hexagonal network of actin is present, sry-α forms a colinear array (Fig. 5A,B). Like actin, the sry-α array is
composed of ‘lines’ of sry-α staining with each line forming
the side of a hexagon. However, the lines of sry-α protein often
do not meet at the hexagon vertices and sry-α is relatively
depleted there. This staining pattern is in marked contrast to
the dots of nullo staining that compose the sides and, most
prominently, the vertices of the hexagons. During early cellularization stages (Fig. 5C,D), sry-α appears to be associated
with the entire plasma membrane. The sry-α protein shows an
additional distinctive localization that is not shared by actin,
myosin or nullo: the protein is localized above each cycle 14
nucleus in a small spherical structure of unknown origin or significance (Fig. 5G,K). This sry-α staining may be associated
with the centrosomes or the microtubule arrays located apical
to the nuclei at these stages (Whitfield et al., 1988; Kellogg et
al., 1989).
Several basic features of sry-α localization are maintained
throughout the slow phase (Fig. 5I,J,K,L,M,N,O,P) and much
of the fast phase (Fig. 5Q,R,S,T) of cellularization. The sry-α
protein remains associated with the invaginating actin-myosin
hexagonal array until late in the fast phase. Although the actinmyosin network assumes a ring-like appearance during fast
phase, sry-α protein continues to form interlocking hexagons
that are composed of lines of sry-α staining (compare Fig. 5M
and 5N). In this regard, sry-α resembles nullo, since neither
protein co-localizes with the rings. It should be noted,
however, that sry-α protein remains localized to the furrow
canal during more advanced stages of cellularization than nullo
does. During cellularization, the association of sry-α with the
plasma membrane is limited to the furrow canal and the lateral
membranes just apical to it. In addition, sry-α protein continues
to be localized to a spherical structure above each nucleus. No
sry-α protein seems to be specifically localized to the apical
plasma membrane, but the protein is present in the cytoplasm
above and, to an increasing extent, below the nuclei. Towards
the end of the fast phase of cellularization (Fig. 5S,T), the
localizations of sry-α protein to the cellularization front and to
the spherical structures above the nuclei are lost and most of
the sry-α protein becomes dispersed throughout the cytoplasm.
The intensity of sry-α antibody staining diminishes rapidly
during the final stages of cellularization, although sry-α protein
remains detectable in gastrulating embryos (data not shown)
until the onset of germ band extension.
Localizations of nullo and sry-α in mutant embryos
The similarities in their mutant phenotypes suggest that nullo
and sry-α proteins may be components of the same develop-
mental pathway. Previous experiments have shown that the
genes are transcriptionally independent (Rose and Wieschaus,
1992). To determine if either protein regulates the translation
or intracellular localization of the other, embryos derived from
either nullo or sry-α stocks were stained with antibodies to the
reciprocal protein.
By approximately 10 minutes into cycle 14, both nullo and
sry-α mutant embryos can be distinguished from sibling
embryos: they exhibit an abnormal array of actin and myosin
that has unusually thin and thick portions and some interfaces
that are completely disrupted. All cycle 14 embryos lacking
sry-α clearly contain nullo protein. The protein forms a
roughly hexagonal network in sry-α mutant embryos with
some disrupted interfaces (Fig. 6M,N,O,P). The discontinuities
in nullo staining completely match the disruptions seen in the
actin-myosin network. Therefore, nullo protein continues to be
coincident with actin and myosin in the sry-α mutant background. Some sry-α embryos appear to have fewer ‘dots’ of
nullo staining than wild-type embryos. It is not clear whether
this decreased intensity of staining is due to an overall decrease
in levels of nullo protein or a slightly compromised ability of
the protein to be properly localized. Overall, however, the distribution of nullo protein is remarkably normal in sry-α
mutants.
In contrast, absence of nullo activity has a more striking
effect on the localization of sry-α protein. While phenotypically nullo embryos contain large pools of sry-α protein, its
distribution pattern is altered. sry-α protein fails to co-localize
with the leading edge of the cellularization front and and the
well defined lines of sry-α staining normally associated with
the actin-myosin array are absent (Fig. 7M,O). Instead, the
entire cytoplasmic region between the nuclei shows a low level
of sry-α staining. Localization of sry-α protein to the spherical
structure above each nucleus persists, however, suggesting that
nullo activity is specifically required for association of sry-α
protein with the actin-myosin network.
DISCUSSION
Translation and post-translational modification of
the nullo protein
Transcription of nullo RNA was previously demonstrated to be
very tightly regulated (Rose and Wieschaus, 1992). The nullo
RNA is expressed only for a brief period during the Drosophila
life cycle: developmental Northern blots revealed the presence
of the nullo transcript in RNA samples from 0- to 4-hour
embryos but not in any other developmental stages. In RNA in
situs to whole mount embryos, the transcript was detectable
from the beginning of cycle 11 through the slow phase of cellularization. Within this short interval, large amounts of nullo
transcript rapidly accumulate, reaching a maximum level
during the division between cycle 13 and 14. As soon as cycle
14 begins, levels of nullo RNA plummet. While accumulation
of the nullo transcript occurs uniformly throughout the embryo,
nullo degradation does not; a reproducibly banded pattern of
the nullo transcript is visible in early cycle 14.
Both developmental western blots of individual, precisely
staged embryos and antibody staining of whole-mount
embryos indicate that the dynamics of nullo protein expression
1872 M. A. Postner and E. F. Wieschaus
mirror those of its RNA. By western analysis, the nullo protein
is detectable from the start of cycle 13 through the beginning
of gastrulation. The highest levels of nullo protein are detected
in embryos in early cycle 14, just prior to and during the slow
phase of cellularization. The levels of nullo protein then drop
rapidly during the fast phase of cellularization. The protein
expression pattern thus closely resembles the transcription
pattern with a translational lag of no greater than twenty
minutes. This comparison implies that nullo protein has a consistently short half-life: nullo protein is constantly being
degraded and replaced with newly synthesized protein until
lack of transcript prevents its replacement. An additional
mechanism of specific degradation late in cellularization
cannot be ruled out.
Independent monoclonal antibodies specifically recognize
two forms of the nullo protein in crude protein preparations
from early embryos. Preliminary results suggest that the size
difference between the two forms reflects a differential phosphorylation, since treatment with bacterial alkaline phosphatase causes the conversion of the slower migrating form
into the faster migrating form (Postner, 1993; and unpublished
observations). The significance of this modification is unclear.
We did not detect pronounced differences in the ratio of phosphorylated to unphosphorylated nullo protein during the entire
window of nullo protein expression.
The nullo and sry-α proteins are components of the
hexagonal network
During early cycle 14, nullo and sry-α proteins both colocalize to the hexagonal array of actin and myosin. They maintain
this association at least until the fast phase of cellularization
(when the cleavage furrow has invaginated about half its final
depth). A more thorough biochemical analysis is required to
determine whether either protein interacts directly with the
actin cytoskeketon. The distributions of the two proteins within
the network are not identical. The sry-α protein is found in
lines running along the sides of the hexagons. These lines
rarely reach the hexagonal vertices and little sry-α protein is
detectable within the vertices. This contrasts with the distribution of nullo protein, which shows a punctate pattern throughout the hexagonal array. nullo appears at regular intervals
along the sides of the hexagons and is particularly abundant at
the hexagonal vertices. The punctate staining pattern is
observed from the earliest stages when the protein can be
detected and suggests a vesicular localization at least during
synthesis or transport to the surface. The potential myristoylation codon at the amino terminus of the nullo protein (Rose
and Wieschaus, 1992) might provide a mechanism for associating the protein with transport vesicles and ultimately with the
plasma membrane. The discovery that the Drosophila virilis
homologue of the nullo protein contains an equally favorable
potential myristoylation codon amidst an otherwise divergent
amino terminus (E. Schejter, personal communication)
strongly suggests that this sequence is important functionally.
If the nullo protein is myristoylated in vivo, nullo might also
provide a crucial link between the actin hexagonal network and
the plasma membrane.
During interphase of cycle 13, nullo forms a network of
interconnecting rings of protein that is associated with the
surface regions between the actin caps. This hexagonal pattern
precedes formation of the actin hexagonal array and might
therefore provide a spatial cue, localizing other proteins into a
hexagonal array following the 13th mitosis. Since an actinmyosin network, albeit abnormal, still forms in nullo mutants,
nullo protein cannot provide the only bias or scaffold for
recruitment of microfilaments and myosin into the hexagonal
pattern. However, other components of the array might be
completely reliant on cycle 13 nullo localization for their
proper distributions; lack of such proteins might cause the
hexagonal array that forms in cycle 14 to be unstable and result
in the disruptions seen in embryos lacking the nullo gene.
Given the similarities between the nullo and sry-α phenotypes,
an obvious possibility is the sry-α protein.
In cellularizing embryos mutant for the nullo locus, the sryα protein is localized to a sphere above each nucleus and distributed throughout the apical cytoplasm just as in wild-type
embryos. However, the protein specifically fails to form a
discrete hexagonal network. Instead, sry-α protein shows a
cytoplasmic localization that is indistinguishable from that
seen in more apical regions. The failure of sry-α protein to
form even a disorganized hexagonal array in embryos lacking
the nullo gene indicates that sry-α is dependent upon nullo for
its localization to the hexagonal network. On the other hand,
nullo protein still co-localizes with the actin network in sry-α
mutants, even though that network is disrupted. This suggests
that the relationship between the loci is not reciprocal and that
sry-α protein is functionally downstream of nullo. The relatively minor effect of sry-α protein on final nullo distribution
is probably indirect: a result of the instability of the actinmyosin network in sry-α mutants.
In this model the nullo phenotype would result in part from
the failure of sry-α to localize to the network and maintain its
stability. This proposal does not account for the difference in
severity between the two null phenotypes. Since more disruptions of the hexagonal network occur in nullo mutant embryos
than in sry-α mutant embryos, the nullo phenotype cannot be
due in its entirety to a mislocalization of sry-α protein. In
addition to facilitating the proper localization of the sry-α
protein, the nullo protein must further contribute to the stability
of the hexagonal array. It could do so indirectly by recruiting
other stabilizing molecules to the network or directly by
binding, and thus anchoring, multiple components of the
network.
We thank Dan Kiehart for the gifts of antibodies and Marty Marlow
of the Princeton’s monoclonal facility for establishing the antibody
lines. Joe Goodhouse, Mark Peifer and Romy Knittel provided
valuable technical advice on confocal microscopy, fusion proteins and
monoclonal antibody screening, respectively. We are indebted to
Alain Vincent for supplying the construct from which the truncated
sry-α protein was produced and for sharing unpublished data and
polyclonal antisera to sry-α with us. This work was supported by grant
5RO1 HD15587 from the National Institutes of Health.
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(Received 4 February 1994 - Accepted 11 April 1994)