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First published online April 10, 2009
doi: 10.1242/10.1242/dev.032425
1 Department of Systems Biology, Harvard Medical School, Boston, MA 02115,
USA.
2 F. M. Kirby Neurobiology Center, Children's Hospital Boston, and Department of
Neurobiology, Harvard Medical School, Boston, MA 02115, USA.
3 Proteomics Center at Children's Hospital Boston, Boston, MA 02115, USA.
4 Department of Pathology, Harvard Medical School and Children's Hospital
Boston, Boston, MA 02115, USA.
* Author for correspondence (e-mail: marc{at}hms.harvard.edu)
Accepted 2 March 2009
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: Yolk, Cell size, Seryp, Vitellogenin, EP45, pNiXa, Xenopus
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Yolk protein is the predominant source of essential amino acids for the
embryo and provides many additional nutrients, including phospholipids,
cholesterol and phosphate. Yolk is stored in dense, membrane-bound organelles
called yolk platelets (YPs). These organelles appear to be highly conserved,
as the YPs of animals in diverse phyla are formed by essentially the same
mechanism and contain homologous proteins
(Fischer et al., 1996
;
Grant and Hirsh, 1999;
Hayakawa et al., 2006;
Opresko et al., 1980;
Roth and Porter, 1964). Yolk
is consumed over the course of embryogenesis, but animals have evolved
different ways of consuming it. The two main patterns, which need not be
mutually exclusive, are: (1) organisms in which these YPs are distributed to
all embryonic cells, which consume the yolk intracellularly (e.g. amphibians,
sea urchins, the nematode Caenorhabditis elegans) and (2) organisms
that develop a yolk sac to digest the YPs and the surrounding cytoplasm and to
distribute nutrients to the embryo proper (e.g. birds, teleost fish, octopi,
insects).
With a few notable exceptions (e.g. sea urchins, dipteran flies), the major
yolk proteins in animals are derived from the large conserved lipoprotein
Vitellogenin (Brooks and Wessel,
2002; Smolenaars et al.,
2007). Vitellogenin is apparently never synthesized by the oocyte,
but is extracted from maternal fluids upon binding to the highly conserved
Vitellogenin receptor (Schneider,
1996). The maternal cell type that provides Vitellogenin differs
in different phyla. Examples include the intestine in C. elegans
(Kimble and Sharrock, 1983),
the fat body in mosquitoes (Hagedorn et
al., 1973) and the liver in vertebrates
(Wahli et al., 1981). Once
endocytosed, Vitellogenin is processed, often being partially proteolyzed into
smaller proteins termed Vitellins or Lipovitellins and, in vertebrates,
Phosvitins and Phosvettes (Romano et al.,
2004). YPs are thought to be specialized late endosomes or
lysosomes, that accumulate and then store the processed Vitellogenin
(Fagotto, 1995;
Grant and Hirsh, 1999).
In X. laevis, YPs occupy around half the volume of the egg, yet
Vitellogenin derivatives account for 
90% (by weight) of egg protein
(Gurdon and Wakefield, 1986).
Such a density of Vitellogenin derivatives is possible because these proteins
are stored in crystalline form within the YP. Vitellogenin derivatives largely
suffice to feed the embryo, probably because Vitellogenin is highly
nutritious. In addition to providing the essential amino acids within its
1800 amino acid chain, Vitellogenin associates with 50 lipid
molecules, 95 molecules of covalently bound phosphate, as well as about
one zinc, two calcium, three magnesium, 0.5 iron and numerous sodium and
potassium ions (Montorzi et al.,
1995; Ohlendorf et al.,
1977; Thompson and Banaszak,
2002; Wiley and Wallace,
1981).
We have gained fundamental insights into the mechanism of YP consumption, a poorly understood process that is likely to be conserved throughout the animal phyla. Our studies have further revealed that yolk consumption is an early aspect of differentiation and that there is a spatial and temporal pattern to nutrition in the Xenopus embryo.
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MATERIALS AND METHODS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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;
Sive et al., 2000). Unless
otherwise specified, chemicals and enzymes were from Sigma-Aldrich (St Louis,
MO, USA).
Purification of yolk platelets and proteomic analysis
Several hundred eggs were activated with the calcium ionophore A23187,
resuspended in YP isolation buffer [YPIB: 20 mM HEPES-KOH pH 7.4, 50 mM KCl,
250 mM sucrose, 1 mM EDTA, 1 mM DTT, 1x Complete protease inhibitors
(Roche, Basel, Switzerland), 100 µg/mL PMSF, 1 µM pepstatin], then lysed
in a loose-fitting Dounce homogenizer. Lysate was layered onto a preformed
Percoll gradient (
=1.12) and centrifuged (30,000 g, 1
hour, 4°C). White, high-density (
1.15) bands were isolated and
the high-density Percoll/organelle mixture was gently mixed and centrifuged
again (20,000 g, 4°C, 30 minutes) to yield two distinct
white bands of YPs. To obtain YPs from tailbud embryos, several hundred
embryos (stage 26) were placed in NKG dissociation buffer [100 mM sodium
isethionate, 20 mM sodium pyrophosphate, 20 mM glucose, pH 9.0
(Newport and Kirschner, 1982)]
with 0.05% benzocaine on a nutator at 4°C for 45 minutes. Single cells or
small cell clusters were passed through a 70 µm cell strainer and were
gently lysed in a loose-fitting Dounce homogenizer. YPs were then purified as
above, but only a single density of YPs was observed. The bands of low-density
egg YPs, high-density egg YPs and tailbud YPs were isolated and
ultracentrifuged (130,000 g, 4°C, 1 hour). Purified YPs
were isolated from the top of the clear Percoll pellets and most were
sonicated. These YP extracts were centrifuged (12,000 g,
4°C, 10 minutes) to pellet YP crystals. This step removed >99% of the
Vitellogenin derivatives Lipovitellin 1 (LV1) and Lipovitellin 2 (LV2). The
12,000 g supernatants were then ultracentrifuged (200,000
g, 4°C, 1 hour) to pellet YP membranes (and any
contaminating vesicles and organelles). The 200,000 g
supernatants were isolated and concentrated in 10 kDa MWCO Centricon tubes
(Millipore, Billerica, MA, USA).
The fractions were subjected to SDS-PAGE followed by staining with
colloidal Coomassie (Invitrogen, Carlsbad, CA, USA). Bands of interest were
excised and in-gel digested with trypsin. Peptides derived from in-gel
digested proteins were analyzed by online microscale capillary reversed-phase
HPLC hyphenated to a linear ion trap mass spectrometer (LTQ, Thermo Fisher
Scientific, Waltham, MA, USA). Samples were loaded onto an in-house packed 100
µm i.d. x 15 cm C18 column (Magic C18, 5 µm,
200 Å, Michrom Bioresources, Auburn, CA, USA) and separated at 500
nL/minute with 30-minute linear gradients from 5 to 40% acetonitrile in 0.4%
formic acid. After each survey spectrum the six most intense ions per cycle
were selected for fragmentation/sequencing. All MS datasets were searched
against a combined X. laevis and X. tropicalis RefSeq
protein sequence database (September 2005, 16994 sequence entries) using the
Mascot search engine (Matrix Science, v. 2.1.04, Boston, MA, USA). Peptides
were identified with a Mascot score no less than 33 (P<0.05) and
proteins were identified based on at least two unique peptides. Pro-Q
Phosphoprotein Stain (Invitrogen) was detected with a Typhoon Trio instrument
(GE Healthcare, Chalfont St Giles, UK). Coomassie staining and western
blotting were imaged with an Odyssey Infrared Imager (Licor, Lincoln, NE,
USA). The same western blot was sequentially probed with different primary
antibodies, but the same fluorescent secondary antibody (goat
anti-rabbit-Alexa680, Invitrogen A21109) was used for each round of
detection.
Antibodies, sectioning and immunostaining
Polyclonal antibodies from rabbit were prepared (Covance, Princeton, NJ,
USA) against Vitellogenin derivatives (anti-Vtg) and against Seryp
(anti-Seryp). The immunogen for anti-Vtg was YP crystals (see above). The
immunogen for anti-Seryp was purified inclusion bodies of untagged Seryp from
Escherichia coli. IgG antibodies were purified from each rabbit serum
and labeled with fluorophores (Invitrogen). Embryos at different stages were
fixed in freshly prepared MEMFA (Sive et
al., 2000), dehydrated and embedded in Paraplast. Sections (10
µm) were rehydrated, microwaved to promote antigen recovery, probed with
anti-Vtg-Alexa488 and anti-Seryp-Alexa568, and z-series separated by
0.3 µm in 30 steps obtained with confocal microscopy using a
100x Plan Apo NA 1.4 objective lens, TE2000U inverted microscope (Nikon,
Melville, NY, USA), CSU10 spinning disk (Yokogawa, Tokyo, Japan), 3W Innova
70C Spectrum laser (Coherent, Santa Clara, CA, USA) and Orca-ER CCD camera
(Hamamatsu, Bridgewater, NJ, USA). For quantitation, YPs were defined as 1
µm bodies, the periphery of which was stained with anti-Vtg. The percentage
of YPs that completely lacked Seryp staining (Seryp-) was
determined. Each tissue at each stage was measured in at least three embryos
derived from different females. The number of YPs counted depended on the size
of the tissue and the YP density, but all the YPs in a selected region were
counted. The number of counts ranged from 41 to 704 YPs/tissue, with a median
of 134 YPs/tissue/embryo.
Dextran injection and tissue dissection
Late blastulae (stage 9) were twice injected in the blastocoel with 10 nL
of 10 mg/mL fixable dextran-Alexa647 (10 kDa, Invitrogen). After developing to
tailbud (stage 26), embryos were transferred to an agarose-coated dish
containing 1xMMR with 0.05% benzocaine and 1 mg/mL collagenase.
Extirpated tissues were transferred into NKG dissociation buffer (see above)
in agarose-coated dishes. Dissociated cells were allowed to settle onto
poly-D-lysine-coated coverslips for 15-45 minutes before being
fixed in freshly prepared MEMFA and processed for immunostaining with
anti-Vtg-Alexa488 and anti-Seryp-Alexa568.
Hydroxyurea/aphidicolin (HU/APH) treatment and CycA2/myc6Cdk2 overexpression
Embryos just beginning gastrulation (stage 10-10.5) were incubated with 20
mM hydroxyurea (HU)/150 µM aphidicolin (APH)/0.5% DMSO or with 0.5% DMSO.
After developing at 22°C for 11.5-12 hours (stage 17-20), embryos were
fixed in MEMFA and the percentage of Seryp- YPs (as above) and the
nuclear density (and hence average cell volume) in the prosencephalon
(n=9 embryos from multiple females) were determined from sagittal
sections. For 6/9 embryos, neighboring sagittal sections from the same embryo
(within 200 µm) were stained with anti-phospho-Histone H3 (serine 10)
(Millipore, 06-570), goat anti-rabbit-Alexa568 (Invitrogen, A11011) and DAPI
to determine the mitotic index in the prosencephalon. Approximately 600 nuclei
were counted (always more than 200) from up to six sections from the same
embryo. To avoid double counting of nuclei, every other 10 µm section was
quantitated. Capped transcripts encoding X. laevis Cyclin A2 (CycA2)
and Cdk2 (with an N-terminal 6myc tag, myc6Cdk2) were prepared with
mMESSAGE Machine (Ambion/Applied Biosystems, Austin, TX, USA). For each
transcript, 250 pg was injected into the animal pole of both blastomeres at
the two-cell stage. Injection of these transcripts had variable,
clutch-dependent effects, including widespread embryo death and gastrulation
defects. Presented data are derived from clutches in which the majority of
injected embryos developed to late neurula (stages 18-19). Fixed embryos
(n=8 embryos from multiple females) were sectioned such that
neighboring transverse sections (always within 200 µm) from the same embryo
could be probed to determine the percentage of Seryp- YPs, nuclear
density and the mitotic index (as above). Anti-myc tag antibody (9E10, Santa
Cruz Biotechnology, Santa Cruz, CA, USA, sc-40) and either goat
anti-mouse-Alexa488 or goat anti-mouse-Alexa647 (Invitrogen A11001, A31625)
identified cells expressing myc6Cdk2 for quantification.
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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),
different embryonic lineages contain different numbers and sizes of YPs.
Therefore, monitoring YP consumption in different embryonic tissues, by
tracking YP numbers or size distributions over time, seemed insensitive and
impractical (Robertson, 1978).
However, we reasoned that the structure of the YP might allow us to
distinguish dormant YPs from those that were engaged in active proteolysis. As
in many amphibians and basal vertebrates, the YPs of X. laevis are
composed of a limiting membrane, a central crystal of Vitellogenin derivatives
and an intervening superficial layer of unknown composition
(Karasaki, 1963;
Romano et al., 2004).
Therefore, if YPs were converted into active degradation compartments during
development (Fagotto, 1995),
components of the superficial layer would be degraded to completion prior to
the crystalline core, as proteases would be likely to only have access to the
outer surface of the protein crystal. Electron microscopy (EM) has revealed
that YPs lacking a superficial layer become increasingly abundant during
amphibian development (Karasaki,
1963).
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;
Wiley and Wallace, 1981).
During oogenesis, endocytosed Vitellogenin is partially proteolyzed into
115 kDa Lipovitellin 1 (LV1), 31 kDa Lipovitellin 2 (LV2), 35
kDa Phosvitin (PHO), 19 kDa Phosvette 1 (PVT1) and 13 kDa Phosvette
2 (PVT2). PVT1 and PVT2 approximate to a cleaved PHO protein and may be
derived from a particular isoform of Vitellogenin
(Ohlendorf et al., 1977;
Wahli et al., 1981;
Wiley and Wallace, 1981). YPs
were purified from frog eggs by isopycnic centrifugation, taking advantage of
the very high density of the organelle. The purified YPs were disrupted by
sonication, and differential centrifugation was used to separate YP crystals
and other insoluble protein from the soluble proteins in the superficial layer
(YP supernatant fraction) and proteins in the limiting membrane (YP membrane
fraction). Our purification scheme removed most Vitellogenin derivatives,
which allowed other YP proteins to be visualized and then identified by
peptide sequencing in a mass spectrometer (see Fig. S1 and Table S1 in the
supplementary material).
A number of plasma and lysosomal proteins were identified. Of interest, we
identified the X. laevis homolog of Paraoxonase, a serum protein that
protects the lipids of the low-density lipoprotein (LDL) from oxidation in
mammals (Getz and Reardon,
2004). Intriguingly, the central protein component of LDL is
apoB100, a lipoprotein that is evolutionarily related to Vitellogenin
(Smolenaars et al., 2007).
Amongst the lysosomal proteins, we identified two different aminopeptidases,
as well as three hydrolases anticipated to attack glycosyl chains. No
additional lysosomal proteases (e.g. Cathepsin D) were found, even though many
of these proteases were represented in the queried sequence databases.
Numerous proteins from the endoplasmic reticulum were identified, in
particular protein disulfide isomerases, as well as many mitochondrial
proteins. Additional tests are required to determine whether these proteins
represent contaminants or bona fide YP proteins.
Aside from Vitellogenin, the most frequently identified peptides originated
from a protein that we have named Seryp (serpin in the yolk platelet).
Previously named EP45 and pNiXa, Seryp is a female-specific serum protein
that, like Vitellogenin, is induced by estrogen and secreted from the liver in
X. laevis (Beck et al.,
1992; Holland et al.,
1992). Seryp is a member of the antitrypsin-like subfamily of the
serpin superfamily (Irving et al.,
2000). Consistently, Seryp has been shown to covalently complex
with, and non-competitively inhibit, bovine chymotrypsin in vitro
(Kotyza et al., 1998).
Although identified in oocytes and embryos, its subcellular location has never
been made clear (Sunderman et al.,
1996). BLAST searches reveal that Seryp is not a well-conserved
protein, being most closely related to two serpins from X. tropicalis
(44% identity over more than 370 amino acids) and to another serpin from
X. laevis (39% identity over more than 360 amino acids) that was not
identified in our YP proteomic study. It is therefore unclear whether Seryp
has been recently recruited to the YP in the X. laevis lineage or if
antitrypsin-like serpins are conserved YP components.
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;
Neff et al., 1984). Anti-Vtg
identified LV1 and LV2, but could not detect the highly phosphorylated PHO,
PVT1 and PVT2. These latter proteins were visualized to varying degrees with
either a Coomassie or a phosphoprotein stain
(Fig. 1A,B). Anti-Seryp
identified three species (p50, p48, p30) that were greatly enriched in the
supernatant fraction of purified YPs that had been disrupted by sonication.
p50 is the most abundant form and has the predicted molecular weight of Seryp;
p48 and p30 are presumably products of partial proteolysis. By this
biochemical analysis, Seryp and PVT1 appear to be the most abundant soluble
proteins in the YP and, as such, are presumably located in the YP superficial
layer. To confirm the localization of Seryp to the YP superficial layer, sections were cut from unfertilized eggs and immunostained (Fig. 1C). Anti-Vtg and anti-Seryp colocalized to YP rims (98%, n=3 eggs, >1450 YPs). Anti-Vtg antibodies apparently could not penetrate the crystalline YP core. But sectioning frequently cracked YPs, exposing the interior of the YP crystal to antibodies. In such cracked YPs, anti-Vtg invariably stained the natural edge of the YP as well as the cracked edge of the crystal (Fig. 1D). By contrast, anti-Seryp only stained the natural edge of the YP, confirming that Seryp is not found within the crystalline core of the YP and is limited to the superficial layer.
Tailbud embryo YPs and the lower density egg YPs mostly lacked proteins in
the YP supernatant fraction (Seryp, PVT1), but did contain proteins from the
YP crystal fraction (LV1, LV2, PHO) (Fig.
1A,B). Curiously, nearly all egg YPs were Vtg+ and
Seryp+ by immunostaining, and subsequent experiments revealed that
many YPs in tailbud embryos still contain Seryp (see below). To determine if
lower density YPs might be derived from a subset of eggs, single eggs were
gently lysed and subjected to isopycnic density fractionation. Single eggs had
lower and higher density YPs (Fig.
1E). Seryp was present in the denser YPs, but it was also found at
the very top of the density gradient where soluble, cytoplasmic proteins would
be found. Immunostaining never revealed substantial Seryp outside of YPs
(Fig. 1C). These observations
can be reconciled by concluding that some aspect of the lysis or density
gradient centrifugation causes a substantial number of egg YPs and all tailbud
YPs to lose their limiting membrane
(Wallace and Karasaki, 1963).
During development, YPs may become closely associated with cytoskeletal
elements (Selchow and Winklbauer,
1997), which might tear the YP membrane during lysis of the
embryonic cells, leading to a complete failure to obtain intact YPs from
tailbud embryos.
Seryp enters the oocyte with Vitellogenin
We wondered whether Seryp, like Vitellogenin, might be a maternal somatic
product that is not synthesized in the oocyte. Given that both Seryp and
Vitellogenin are estrogen-responsive serum proteins that are secreted from the
liver (Holland et al., 1992),
it seemed plausible that Seryp also entered YPs by endocytosis during
oogenesis. We prepared extracts of staged oocytes
(Fig. 2A). Egg forms of Seryp
(p50, p48, p30) were always present in later oocyte stages (III-VI, 15 of 15).
Strikingly, in the earliest stages (I,II), oocytes did not always contain the
standard forms of Seryp but did contain a novel protein (p60) detected by our
anti-Seryp antibody. p60 was present in all stage I and II oocytes (5 of 5)
and in most stage III oocytes (3 of 4). p60 abundance in later stage oocytes
(IV-VI) was more variable (see Fig. S2 in the supplementary material).
In reciprocal co-immunoprecipitation experiments, Seryp and Vitellogenin
interacted in X. laevis plasma
(Fig. 2B), as originally
suggested by ultracentrifugation experiments
(Holland and Wangh, 1987).
Minor yolk proteins in chicken interact with the Vitellogenin receptor,
accounting for their concentration in the yolk
(Jacobsen et al., 1995).
Interaction with Vitellogenin may be an additional mechanism for targeting
proteins to the YP. As Seryp is far less abundant in plasma than Vitellogenin
(more than 50-fold molar difference, data not shown), only a fraction of the
Vitellogenin entering the oocyte need interact with Seryp to account for the
abundance of Seryp in oocytes and eggs.
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1 transcript per typical metazoan cell or
100,000 transcripts per frog oocyte, egg or embryo. Using this number as
the maximal possible amount of SERYP transcript present in oocytes or
eggs, simple calculations show that Seryp translation in oocytes and eggs, if
any, is insignificant relative to the massive accumulation and degradation of
this protein (1 µg per egg) during oogenesis and embryogenesis,
respectively. We conclude that, like Vitellogenin, Seryp is a maternal somatic
protein that is endocytosed and stored by oocytes and then consumed during
embryogenesis.
Seryp is eliminated from embryos and from individual YPs prior to Vitellogenin derivatives
Although the bulk of Vitellogenin derivatives and Seryp were not degraded
until late in embryogenesis, Seryp species clearly declined prior to the
Vitellogenin derivatives in whole embryos
(Fig. 3A,B), as previously
suggested (Grbac-Ivankovic et al.,
1994). Variability in the absolute YP protein content of X.
laevis eggs (Fig. 3B), and
the fact that most of the yolk protein is concentrated in the presumptive gut
(Hausen and Riebesell, 1991),
precluded using bulk measurements of Seryp and Vitellogenin derivatives as a
useful indicator of yolk consumption.
As described above, we suspected that within individual YPs a superficial-layer protein like Seryp would be degraded prior to the central crystal of Vitellogenin derivatives. Indeed, in some tissues of tailbud embryos, many YPs completely lacked Seryp (Seryp-) but did contain Vitellogenin derivatives (Vtg+) (Fig. 3C). As essentially all egg YPs stained positive for Seryp (Fig. 1C), yolk proteolysis had apparently initiated in these tissues.
Yolk platelets that lack Seryp have been reincorporated into the endocytic system in a quantal fashion
There is no consensus on how yolk is proteolyzed intracellularly. In the
most cited current model, it is argued that in X. laevis and many
other species YPs need only be acidified to activate preloaded proteases
(Fagotto, 1995). This model
was based on observations that YPs become more acidic during development in
many organisms and protease activities can be found in dormant YPs. In an
alternative model, YPs would be activated by fusion with endosomes or
lysosomes. The increased acidity of active YPs would be a secondary
consequence of endocytic proton pumps, and not an independent regulatory step.
The entire repertoire of lysosomal hydrolases and transporters would then be
available to hydrolyze the diverse substrates present in the YP (see
Introduction) and to export small molecules. Previously, in newt embryos,
endocytosed ferritin was observed by EM in YPs that appeared to be in the
process of degradation (Komazaki and
Hiruma, 1999).
To test the endosome fusion model, we injected fixable dextran conjugated
to a fluorescent dye into the blastocoel (i.e. extracellular space) of late
blastulae (stage 9). The embryos were reared to tailbud stage (stage 26),
select tissues dissected and cells dissociated. Endocytosed dextran had
accumulated in a subset of YPs in many cells
(Fig. 4A,B). Of the
dextran+ bodies larger than 1 µm (n=349 from 149
cells), 91% were Vtg+ YPs. Strikingly, only 3% of these
dextran-positive bodies were Seryp+. There were still many
Vtg+ Seryp+ YPs present in these cells
(Fig. 4A,B). In short, the
subsets of Vtg+ YPs that were dextran+ and
Seryp+ were nearly mutually exclusive. The data argue that the
subset of YPs that are engaged in proteolysis have also been mixing with
compartments of the endocytic system. We will refer to this subset of YPs as
being `activated'. Since they accumulate most of the dextran present in the
cell (Fig. 4A,B), activated YPs
appear to be the principal terminal degradative compartments in frog embryonic
cells. As single cells from a variety of tissues contained both activated and
dormant YPs (Fig. 4C), YP
proteolysis is, per platelet, a quantal process within embryonic cells.
|
Several tissues in early neurulae were avidly consuming yolk
(Fig. 5C). Although it avoids
the perils of quantitating fluorescence, our binary YP classification has an
inherent delay, namely the time to degrade Seryp to completion. Therefore, it
is not possible to say when YP consumption is first initiated. A previous
histological survey concluded that YP consumption commences much later in
embryogenesis, starting in the tailbud stages
(Selman and Pawsey, 1965). By
contrast, there have been scattered reports of YP breakdown prior to the
tailbud stages in various amphibians
(Fagotto and Maxfield, 1994;
Karasaki, 1963;
Komazaki et al., 2002;
Robertson, 1978). Our results
provide a conclusive demonstration that yolk consumption occurs throughout
nearly the whole of X. laevis embryogenesis.
Activated YPs generally correlated with morphological differentiation.
Ectodermal cells and the dorsal mesodermal cells differentiate in structure
during neurulation, when the percentage of activated YPs abruptly rises. Among
ectodermal tissues, the prosencephalon and eye had the highest percentages of
activated YPs. Induction of the cement gland and lens from epidermis
correlated with increased YP consumption
(Fig. 5A,B). Morphological
changes to the endoderm first occur in tailbud embryos, when the visceral
pouches form during stages 23-27
(Nieuwkoop and Faber, 1994).
Consistently, marked YP consumption took place in the arch and oral endodermal
epithelium of tailbud embryos (stage 27), but not in morphologically static
deep endoderm (Fig. 5A,B).
Most YP protein is, however, inherited by the deep endoderm that develops
into the gut. Degradation of the bulk of Seryp and Vitellogenin derivatives
accompanies gut development during the later tailbud and tadpole stages
(Fig. 3A,B). It has been
thought that the YPs of the gut are degraded extracellularly, by digestion of
yolky cells within the intestinal lumen
(Gerhart, 1980;
Hausen and Riebesell, 1991).
However, recent fate maps suggest that all deep endoderm cells are
incorporated into the intestinal wall
(Chalmers and Slack, 2000).
Cells of the developing intestine wall, in fact, contained numerous
intracellular, activated YPs (see Fig. S4 in the supplementary material).
Therefore, even in the developing gut, yolk consumption was intracellular and
correlated with morphological differentiation.
The onset of yolk consumption is not triggered by small cell size
Our survey of YP activation indicated that something happens between the
blastula and early neurula stages to trigger this process, particularly in the
neurectoderm. In the blastula, animal pole cells, which will give rise to the
neurectoderm, are relatively small owing to previous asymmetric animal-vegetal
divisions. The neurectoderm is also the most actively dividing tissue in early
neurulae (Saka and Smith,
2001). Neurectoderm cells are therefore likely to be the smallest
in the neurula and these cells also exhibit the largest percentage of active
YPs (Fig. 5). We considered the
possibility that small cell size induces compensatory cell growth and hence YP
consumption.
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|
; Harris and
Hartenstein, 1991). HU/APH-treated embryos had oversized cells,
but degraded YPs as rapidly as control siblings
(Fig. 6A,C-E). We performed the
inverse experiment by generating undersized cells
(Richard-Parpaillon et al.,
2004). Cells expressing exogenous CycA2 and myc6Cdk2
(myc+) (Fig. 6B) displayed both
a higher frequency of mitotic cells and a significant decrease in average cell
size (15%) (Fig. 6B-D),
demonstrating that CycA2/myc6Cdk2 was driving cell cycle
progression and not just prolonging mitosis
(den Elzen and Pines, 2001).
Despite excessive cell cycle progression and small cell size,
CycA2/myc6Cdk2-expressing cells were not engaged in more rapid YP
consumption (Fig. 6E). |
DISCUSSION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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), allows extracellular dextran
(Fig. 4) and ferritin access to
the YP (Komazaki and Hiruma,
1999) and leads to the proteolysis of Seryp and, eventually, the
crystal of Vitellogenin derivatives. In addition to proteases, phosphatases
and other hydrolases, fusion events with late endosomes or lysosomes would
supply lipid, amino acid and ion transporters. The quantal mechanism of YP consumption appears to provide cells with a valve to control the rate of nutrient release. If cells did not single out a subset of YPs, but instead activated all YPs simultaneously, several problems would be immediately apparent. The endocytic system would first need to cope with a rapid expansion in volume. Even if rapidly increasing the size of the endocytic system were not a problem, it is not clear how nutrient release would be controlled in such a scenario. To control nutrient release, this hypothetical cell could conceivably regulate the number of hydrolases provided to all of the YPs. By contrast, real embryonic cells appear to have a much simpler regulatory system. YPs are individually activated, perhaps at a rate controlled by intracellular nutrient sensors, and then degraded to completion.
The striking spatial differences in yolk consumption argues that nutrition
in early frog embryos is tissue- and/or cell-autonomous. That is, tissues that
differentiate early, such as the neural lineages and the somites, have a
requirement for components of the YP and they obtain these components
intracellularly, with cells in that tissue perhaps sharing nutrients through
gap junctions. An alternative possibility, which our evidence does not
support, was that all cells in the early embryo would simultaneously begin
degrading yolk and sharing nutrients through the extracellular space. In later
stage tailbud embryos, the developing vasculature may allow for an embryo-wide
integration of nutrition (heart beating begins at stage 33-34). The vegetal
mass (presumptive amphibian gut) inherits most of the yolk protein and yet
comes to represent a proportionally smaller part of the embryo during the
tadpole stages (Nieuwkoop and Faber,
1994). Nutrients released by intracellular breakdown of YPs within
the cells of the gut are presumably exported into the blood and taken up by
growing tissues during these later stages of embryogenesis.
Why do differentiating tissues consume yolk? This question is complicated
by the fact that yolk consumption will release many different substances: not
just amino acids, but cholesterol, phospholipids, fatty acids, phosphate and
numerous positively charged ions. Differentiation per se requires no net
synthesis of proteins: extant proteins can be degraded to allow for the
remodeling of the proteome. But differentiation will certainly require energy,
perhaps more energy than is required to maintain a cell that is not
differentiating. Although it is often thought that yolk is consumed to provide
energy, most evidence from early frog embryos argues against this. Free amino
acids may provide some energy during the cleavage stages
(Dworkin and Dworkin-Rastl,
1991) but, starting at gastrulation, the respiratory quotient and
lack of ammonia production suggest that most respiration is driven by
carbohydrate, and not amino acid, oxidation
(Boell, 1948). Indeed, the frog
egg has extensive energetic stores: 5% of the dry mass of amphibian eggs
is glycogen, and triacylglycerides are packed into abundant lipid droplets
(Billett and Gould, 1971;
Dworkin and Dworkin-Rastl,
1991; Hausen and Riebesell,
1991).
We argue that YP consumption in some tissues could signify a specialized
form of growth, in which stored yolk protein is converted into the functional
biomass of the embryo, such as cytoplasm and extracellular matrix. Growth has
rarely been considered in such terms. Some might reasonably insist that growth
can only be defined as an increase in total mass. But frog embryos clearly
increase in volume (Tuft,
1962) (even when large extracellular spaces like the archentron
are accounted for), synthesize new ribosomes
(Baum and Wormington, 1985;
Pierandrei-Amaldi et al.,
1982) and accumulate large amounts of non-yolk proteins (see Fig.
S5 in the supplementary material). Furthermore, few would dispute that chick
embryos grow at the expense of their extra-embryonic yolk supply. The
intracellular location of the yolk in amphibians (and many other organisms)
does not change the fact that it is essentially a food that provides for
growth of the embryonic tissues. It remains to be determined whether the
canonical growth control pathways of somatic cells, such as the TOR pathway,
control yolk consumption.
Aside from its small size (1 mm) and amenability to biochemical,
chemical, genetic and genomics approaches, the cell-autonomous nature of
nutrition in early X. laevis embryos offers pleasing simplifications.
Small molecule or genetic perturbations that incidentally cripple the nervous
system, gut, or vasculature will block nutrition in feeding animals, but will
not necessarily do so in frog embryonic cells laden with YPs, lipid droplets
and glycogen. Future work in frog embryos might inform many aspects of
metabolism in the embryonic and adult stages of vertebrate life.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/9/1539/DC1
|
ACKNOWLEDGMENTS |
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|
Footnotes |
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|
REFERENCES |
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|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
Baum, E. Z. and Wormington, W. M. (1985).
Coordinate expression of ribosomal protein genes during Xenopus
development. Dev. Biol.
111,488
-498.[CrossRef][Medline]
Beck, B. L., Henjum, D. C., Antonijczuk, K., Zaharia, O., Korza,
G., Ozols, J., Hopfer, S. M., Barber, A. M. and Sunderman, F. W., Jr
(1992). pNiXa, a Ni(2+)-binding protein in Xenopus
oocytes and embryos, shows identity to Ep45, an estrogen-regulated hepatic
serpin. Res. Commun. Chem. Pathol. Pharmacol.
77, 3-16.[Medline]
Billett, F. S. and Gould, R. P. (1971). Fine
structural changes in the differentiating epidermis of Xenopus laevis
embryos. J. Anat. 108,465
-480.[Medline]
Boell, E. J. (1948). Biochemical
differentiation during amphibian development. Ann. NY Acad.
Sci. 49,773
-800.[CrossRef][Medline]
Brooks, J. M. and Wessel, G. M. (2002). The
major yolk protein in sea urchins is a transferrin-like, iron binding protein.
Dev. Biol. 245,1
-12.[CrossRef][Medline]
Chalmers, A. D. and Slack, J. M. (2000). The
Xenopus tadpole gut: fate maps and morphogenetic movements.
Development 127,381
-392.[Abstract]
Danilchik, M. V. and Gerhart, J. C. (1987).
Differentiation of the animal-vegetal axis in Xenopus laevis oocytes.
I. Polarized intracellular translocation of platelets establishes the yolk
gradient. Dev. Biol.
122,101
-112.[CrossRef][Medline]
den Elzen, N. and Pines, J. (2001). Cyclin A is
destroyed in prometaphase and can delay chromosome alignment and anaphase.
J. Cell Biol. 153,121
-136.
Dworkin, M. B. and Dworkin-Rastl, E. (1991).
Carbon metabolism in early amphibian embryos. Trends Biochem.
Sci. 16,229
-234.[CrossRef][Medline]
Fagotto, F. (1995). Regulation of yolk
degradation, or how to make sleepy lysosomes. J. Cell
Sci. 108,3645
-3647.[Medline]
Fagotto, F. and Maxfield, F. R. (1994). Changes
in yolk platelet pH during Xenopus laevis development correlate with
yolk utilization. A quantitative confocal microscopy study. J. Cell
Sci. 107,3325
-3337.[Abstract]
Fischer, A., Dorresteijn, A. W. and Hoeger, U.
(1996). Metabolism of oocyte construction and the generation of
histospecificity in the cleaving egg. Lessons from nereid annelids.
Int. J. Dev. Biol. 40,421
-430.[Medline]
Gerhart, J. C. (1980). Mechanisms regulating
pattern formation in the amphibian egg and early embryo. In
Biological Regulation and Development, Vol. 2, Molecular
Organization and Cell Function (ed. R. F. Goldberger), pp.133
-316. New York: Plenum Press.
Getz, G. S. and Reardon, C. A. (2004).
Paraoxonase, a cardioprotective enzyme: continuing issues. Curr.
Opin. Lipidology 15,261
-267.[CrossRef][Medline]
Grant, B. and Hirsh, D. (1999).
Receptor-mediated endocytosis in the Caenorhabditis elegans oocyte.
Mol. Biol. Cell 10,4311
-4326.
Grbac-Ivankovic, S., Antonijczuk, K., Varghese, A. H., Plowman,
M. C., Antonijczuk, A., Korza, G., Ozols, J. and Sunderman, F. W., Jr
(1994). Lipovitellin 2 beta is the 31 kD Ni(2+)-binding protein
(pNiXb) in Xenopus oocytes and embryos. Mol. Reprod.
Dev. 38,256
-263.[CrossRef][Medline]
Gurdon, J. B. and Wakefield, L. (1986).
Microinjection of amphibian oocytes and eggs for the analysis of
transcription. In Microinjection and Organelle Transplantation
Techniques (ed. J. E. Celis, A. Graessmann and A. Loyter).
London: Academic Press.
Hagedorn, H. H., Fallon, A. M. and Laufer, H.
(1973). Vitellogenin synthesis by the fat body of the mosquito
Aedes aegypti: evidence of transcriptional control. Dev.
Biol. 31,285
-294.[CrossRef][Medline]
Hardcastle, Z. and Papalopulu, N. (2000).
Distinct effects of XBF-1 in regulating the cell cycle inhibitor p27(XIC1) and
imparting a neural fate. Development
127,1303
-1314.[Abstract]
Harris, W. A. and Hartenstein, V. (1991).
Neuronal determination without cell division in Xenopus embryos.
Neuron 6,499
-515.[CrossRef][Medline]
Hausen, P. and Riebesell, M. (1991).
The Early Development of Xenopus laevis: An Atlas of the
Histology. Berlin: Springer-Verlag.
Hayakawa, H., Andoh, T. and Watanabe, T.
(2006). Precursor structure of egg proteins in the coral Galaxea
fascicularis. Biochem. Biophys. Res. Commun.
344,173
-180.[CrossRef][Medline]
Holland, L. J. and Wangh, L. J. (1987).
Estrogen induction of a 45 kDa secreted protein coordinately with vitellogenin
in Xenopus liver. Mol. Cell. Endocrinol.
49, 63-73.[CrossRef][Medline]
Holland, L. J., Suksang, C., Wall, A. A., Roberts, L. R., Moser,
D. R. and Bhattacharya, A. (1992). A major estrogen-regulated
protein secreted from the liver of Xenopus laevis is a member of the
serpin superfamily. Nucleotide sequence of cDNA and hormonal induction of
mRNA. J. Biol. Chem.
267,7053
-7059.
Irving, J. A., Pike, R. N., Lesk, A. M. and Whisstock, J. C.
(2000). Phylogeny of the serpin superfamily: implications of
patterns of amino acid conservation for structure and function.
Genome Res. 10,1845
-1864.
Jacobsen, L., Hermann, M., Vieira, P. M., Schneider, W. J. and
Nimpf, J. (1995). The chicken oocyte receptor for lipoprotein
deposition recognizes alpha 2-macroglobulin. J. Biol.
Chem. 270,6468
-6475.
Jared, D. W., Dumont, J. N. and Wallace, R. A.
(1973). Distribution of incorporated and synthesized protein
among cell fractions of Xenopus oocytes. Dev.
Biol. 35,19
-28.[CrossRef][Medline]
Karasaki, S. (1963). Studies on amphibian yolk.
5. Electron microscopic observations on the utilization of yolk platelets
during embryogenesis. J. Ultrastruct. Res.
59,225
-247.[Medline]
Kimble, J. and Sharrock, W. J. (1983).
Tissue-specific synthesis of yolk proteins in Caenorhabditis elegans.
Dev. Biol. 96,189
-196.[CrossRef][Medline]
Komazaki, S. and Hiruma, T. (1999). Degradation
of yolk platelets in the early amphibian embryo is regulated by fusion with
late endosomes. Dev. Growth Differ.
41,173
-181.[CrossRef][Medline]
Komazaki, S., Tanaka, N. and Nakamura, H.
(2002). Regional differences in yolk platelet degradation
activity and in types of yolk platelets degraded during early amphibian
embryogenesis. Cells Tissues Organs
172, 13-20.[CrossRef][Medline]
Kotyza, J., Varghese, A. H., Korza, G. and Sunderman, F. W.,
Jr (1998). Interactions of serine proteinases with pNiXa, a
serpin of Xenopus oocytes and embryos. Biochim. Biophys.
Acta. 1382,266
-276.[CrossRef][Medline]
Montorzi, M., Falchuk, K. H. and Vallee, B. L.
(1995). Vitellogenin and lipovitellin: zinc proteins of
Xenopus laevis oocytes. Biochemistry
34,10851
-10858.[CrossRef][Medline]
Neff, A. W., Wakahara, M., Jurand, A. and Malacinski, G. M.
(1984). Experimental analyses of cytoplasmic rearrangements which
follow fertilization and accompany symmetrization of inverted Xenopus
eggs. J. Embryol. Exp. Morphol.
80,197
-224.[Medline]
Newport, J. and Kirschner, M. (1982). A major
developmental transition in early Xenopus embryos: I.
Characterization and timing of cellular changes at the midblastula stage.
Cell 30,675
-686.[CrossRef][Medline]
Nieuwkoop, P. D. and Faber, J. (1994).
Normal Table of Xenopus laevis (Daudin), 2nd edn. New
York: Garland Publishing.
Ohlendorf, D. H., Barbarash, G. R., Trout, A., Kent, C. and
Banaszak, L. J. (1977). Lipid and polypeptide components of
the crystalline yolk system from Xenopus laevis. J. Biol.
Chem. 252,7992
-8001.
Opresko, L., Wiley, H. S. and Wallace, R. A.
(1980). Differential postendocytotic compartmentation in
Xenopus oocytes is mediated by a specifically bound ligand.
Cell 22,47
-57.[CrossRef][Medline]
Pierandrei-Amaldi, P., Campioni, N., Beccari, E., Bozzoni, I.
and Amaldi, F. (1982). Expression of ribosomal-protein genes
in Xenopus laevis development. Cell
30,163
-171.[CrossRef][Medline]
Richard-Parpaillon, L., Cosgrove, R. A., Devine, C., Vernon, A.
E. and Philpott, A. (2004). G1/S phase cyclin-dependent
kinase overexpression perturbs early development and delays tissue-specific
differentiation in Xenopus. Development
131,2577
-2586.
Robertson, N. (1978). Labilization of the
superficial layer and reduction in size of yolk platelets during early
development of Xenopus laevis. Cell Differ.
7, 185-192.[CrossRef]
Romano, M., Rosanova, P., Anteo, C. and Limatola, E.
(2004). Vertebrate yolk proteins: a review. Mol.
Reprod. Dev. 69,109
-116.[CrossRef][Medline]
Roth, T. F. and Porter, K. R. (1964). Yolk
protein uptake in the oocyte of the mosquito Aedes aegypti. L.
J. Cell Biol. 20,313
-332.
Saka, Y. and Smith, J. C. (2001). Spatial and
temporal patterns of cell division during early Xenopus
embryogenesis. Dev. Biol.
229,307
-318.[CrossRef][Medline]
Sambrook, J. and Russell, D. W. (2001).
Molecular Cloning: A Laboratory Manual. Cold Spring
Harbor, NY: Cold Spring Harbor Laboratory Press.
Schneider, W. J. (1996). Vitellogenin
receptors: oocyte-specific members of the low-density lipoprotein receptor
supergene family. Int. Rev. Cytol.
166,103
-137.[Medline]
Selchow, A. and Winklbauer, R. (1997).
Structure and cytoskeletal organization of migratory mesoderm cells from the
Xenopus gastrula. Cell Motil. Cytoskeleton
36, 12-29.[CrossRef][Medline]
Selman, G. G. and Pawsey, G. J. (1965). The
utilization of yolk platelets by tissues of Xenopus embryos studied
by a safranin staining method. J. Embryol. Exp.
Morphol. 14,191
-212.[Medline]
Sive, H. L., Grainger, R. M. and Harland, R. M.
(2000). Early Development of Xenopus laevis: A
Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor
Laboratory Press.
Smolenaars, M. M., Madsen, O., Rodenburg, K. W. and Van der
Horst, D. J. (2007). Molecular diversity and evolution of the
large lipid transfer protein superfamily. J. Lipid
Res. 48,489
-502.
Sunderman, F. W., Jr, Varghese, A. H., Kroftova, O. S.,
Grbac-Ivankovic, S., Kotyza, J., Datta, A. K., Davis, M., Bal, W. and
Kasprzak, K. S. (1996). Characterization of pNiXa, a serpin
of Xenopus laevis oocytes and embryos, and its histidine-rich,
Ni(II)-binding domain. Mol. Reprod. Dev.
44,507
-524.[CrossRef][Medline]
Thompson, J. R. and Banaszak, L. J. (2002).
Lipid-protein interactions in lipovitellin.
Biochemistry 41,9398
-9409.[CrossRef][Medline]
Tuft, P. H. (1962). The uptake and distribution
of water in the embryo of Xenopus laevis (Daudin). J. Exp.
Biol. 39,1
-19.
Wahli, W., Dawid, I. B., Ryffel, G. U. and Weber, R.
(1981). Vitellogenesis and the vitellogenin gene family.
Science 212,298
-304.
Wallace, R. A. and Karasaki, S. (1963). Studies
on amphibian yolk. 2. The isolation of yolk platelets from the eggs of
Rana pipiens. J. Cell Biol.
18,153
-166.
Wiley, H. S. and Wallace, R. A. (1981). The
structure of vitellogenin: multiple vitellogenins in Xenopus laevis
give rise to multiple forms of the yolk proteins. J. Biol.
Chem. 256,8626
-8634.
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-Vtg) and anti-Seryp-Alexa568 (
