V-ATPase expression during development of Artemia franciscana embryos: potential role for proton gradients in anoxia signaling
Division of Cellular, Developmental and Integrative Biology, Department of Biological Science, Louisiana State University, Baton Rouge, LA 70803, USA
* Author for correspondence (e-mail: jcovi1{at}lsu.edu)
Accepted 11 May 2005
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Summary |
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Key words: V-type ATPase, mRNA expression, protein expression, quiescence, diapause, brine shrimp, B-subunit cDNA, acidification, bafilomycin, oligomycin, hatching success
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Introduction |
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It is important to note that the ability of this free-floating gastrula to
downregulate metabolism in response to anoxia (Hand and Gnaiger, 1988) is
present from the moment development begins, but is subsequently lost when they
emerge from their protective cyst coat after 812 h of aerobic
development (Ewing and Clegg,
1969; Stocco et al.,
1972
). The encompassing cyst coat, restricted to the embryonic
stage, isolates the embryo from its environment with a highly selective
permeability barrier (for discussion, see
Trotman, 1991
). This barrier
precludes the involvement of exogenous H+ equivalents in the
previously discussed acidification (Busa et
al., 1982
). Unfortunately, the nature of the cyst shell confounds
examination of the proton-generating mechanisms responsible for the pH
transition by preventing the direct measurement of pHi in
vivo with the use of microelectrodes or reporter compounds. For this
reason, 31P-NMR is the only method to yet provide in vivo
measurements of embryonic pHi
(Busa et al., 1982
;
Busa and Nuccitelli, 1984
;
Clegg et al., 1995
;
Kwast et al., 1995
), although
a limitation inherent in this method suggests that acidic compartments may be
involved in anoxia signaling.
While the caveats of using 31P-NMR to determine in vivo
pHi have been covered elsewhere
(Busa et al., 1982), it is
important to note here that the technique is limited by its dependence on the
presence of free inorganic phosphate (Pi) in the compartment(s)
being observed. Consequently, compartments having low Pi
concentration or low relative volume have a negligible effect on the average
pH being measured for a population of whole embryos. In other words, protons
stored in these compartments would remain undetected until their release into
a space having a relatively high Pi content, whereupon the event
would be observed as an acidification. One potentially undetected compartment
might be the extracellular space, which has been noted in some organisms to
contain as little as 20% of the Pi found in the cytoplasm
(Grabe and Oster, 2001
).
Alternatively, undetected acidic compartments may be subcellular in nature,
and their visualization precluded by the fact that they only contain a small
percentage of free embryonic Pi. These presumably would include
Golgi, tubules, exocytotic vesicles, coated vesicles, early and late
endosomes, and lysosomes, all of which are likely to have functional roles
during such events as production of the embryonic cuticle (cf.
Criel, 1991
), formation of
yolk platelets (Giorgi et al.,
1999
; Warner et al.,
2002
) and yolk degradation
(Komazaki and Hiruma, 1999
;
Perona et al., 1988
; Perona
and Vallejo, 1985
,
1989
). Indeed, membrane-bound
yolk platelets themselves should be considered, as they are acidified to
varying degrees for both maintenance and degradation in other species
(Abreu et al., 2004
;
Fagotto, 1995
;
Fausto et al., 2001
) and
contain very little Pi in A. franciscana
(Warner and Huang, 1979
). An
interesting commonality among all of these compartments is that their final
luminal pH is set in part by the V-ATPase proton pump
(Futai et al., 1998
;
Grabe and Oster, 2001
;
Nishi and Forgac, 2002
), and
thus this enzyme became the focal point of our research.
The V-ATPase is a multimeric complex composed of two primary domains
(V1 and V0). The extra-membrane V1 domain is
comprised of eight different subunits, labeled AH, and is responsible
for ATP hydrolysis (Wieczorek et al.,
2000). The membrane-spanning V0 domain consists of at
least three different subunits, labeled a, d and c, and is responsible for
providing a proton path across the membrane
(Forgac, 1998
). When combined,
these two domains form a molecular motor well known for both its broad
distribution and diverse functional capacity for proton transport. For
example, in addition to their role in acidification of intracellular
compartments (Forgac, 2000
;
Futai et al., 2000
), V-ATPases
are also known to power acid secretion and secondary active transport in
animal plasma membranes (for recent reviews, see
Kawasaki-Nishi et al., 2003
;
Nelson and Harvey, 1999
;
Nelson et al., 2000
;
Wieczorek et al., 2000
).
Taking these facts into consideration, the V-ATPase is a prime candidate for
facilitating the proton transport required to establish intracellular and/or
extracellular acidic compartments in A. franciscana embryos.
We hypothesize that, in the face of severe energetic constraints imposed by anoxia, H+ gradients between acidic compartments and the surrounding cytoplasm dissipate, producing an acidification critical to metabolic downregulation in A. franciscana embryos. As first steps in addressing this hypothesis, we examined expression of the V-ATPase during the early development of these embryos and demonstrate that in vivo inhibition of this proton pump by bafilomycin blocks normoxic development.
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Materials and methods |
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Manduca sexta were obtained as 3rd-instar larvae from a colony at
the Arizona Research Labs Division of Neurobiology, University of Arizona
(Willis and Arbas, 1998).
Dissections were performed according to Harvey et al.
(1990
) after larvae developed
to the feeding phase of the 4th-instar stage.
Molecular cloning and sequencing of cDNA for subunit B of the V-ATPase
AMV reverse transcriptase (Promega, Madison, WI, USA) and a d18TV primer
were employed to reverse transcribe poly(A+) mRNA from DNase-treated total
RNA, previously isolated from 6 h aerobic A. franciscana embryos
(Hardewig et al., 1996). A
central domain of the V-ATPase B-subunit was amplified using degenerate
primers (HATF2, HATR4; Weihrauch et al.,
2001
; Table 1).
Following agarose gel purification, the amplified cDNA was cloned, and the
insert was sequenced directly from the plasmid construct. After the identity
of the cDNA fragment was confirmed by BLAST search of the GenBank database,
the 3' and 5' ends were amplified by rapid amplification of cDNA
ends (RACE) using a First Choice RLM RACE kit (Ambion, Austin, TX, USA)
according to the manufacturer's instructions. The 5' end was amplified
from total RNA isolated from post-diapause embryos immediately after hydration
(hour 0). Since amplification of the 3' end is more efficient when mRNA
is used, the 3' end was amplified from mRNA previously isolated from
diapause embryos. It is important to note that, because diapause embryos are
developmentally arrested, the mRNA of these embryos is theoretically
equivalent to that of post-diapause embryos before development is resumed
(hour 0). RACE PCR products were gel purified, cloned and sequenced directly
from the plasmid construct using specific primers designed with the assistance
of Primer3 software (Rozen and Skaletsky,
2000
). Sequencing utilized BigDye terminator chemistry and an ABI
PRISM 3100 Genetic Analyzer (Applied Biosystems, Foster City, CA, USA).
Sequences were assembled using Sequencher software (Gene Codes Co., Ann Arbor,
MI, USA). An open reading frame (ORF) was identified using a freely available
ORF tool available from The National Center for Biotechnology Information
(http://www.ncbi.nlm.nih.gov).
DNA-to-protein translation, pI estimation and theoretical calculation of
relative molecular mass (Mr) were achieved using tools
made available by the Swiss Institute of Bioinformatics
(http://us.expasy.org/tools).
Multiple sequence alignments were produced with Clustal X version 1.83
(Thompson et al., 1997
).
Sequences were annotated with GeneDoc version 2.6.002
(http://www.psc.edu/biomed/genedoc).
Phylogenetic analysis was conducted with MEGA version 2.1
(Kumar et al., 2004
). Based on
the work of Hillis and Bull
(1993
), all bootstrap
proportions
70% were considered strong support for a given clade.
|
Isolation of total RNA
RNase-free conditions were maintained throughout all isolations using
published methods (Hardewig et al.,
1996). Encysted embryos were washed using a modified
dechorionation protocol (Kwast and Hand,
1993
). All wash solutions were maintained at 0°C to prevent
premature development and complete removal of the chorion. Aseptic embryos
were then incubated at 2223°C with shaking at 110 r.p.m. in
Erlenmeyer flasks containing 35
artificial seawater.
Two different procedures were employed for isolating total RNA. In the
first case, hydrated embryos were kept on ice until the start of incubation,
and starting times were staggered, allowing all time points to be processed
within 20 min of each other. Developing embryos were equilibrated with a 40:60
O2:N2 gas mixture (cf.
Carpenter and Hand, 1986), and
triplicate extractions were performed for each time point by homogenization in
a guanidinium thiocyanate-based buffer, followed by isolation of total RNA by
centrifugation through a CsCl cushion as described by Hardewig et al.
(1996
). Alternatively,
incubation of hydrated embryos was initiated synchronously, and development
proceeded in a medium equilibrated with room air. At predetermined intervals,
triplicate flasks of embryos were suspended in ice-cold 35
artificial
seawater to arrest development until homogenization. For each time point,
embryos were filtered, rinsed with cold DEPC water and blotted dry. RNA
extractions were performed with an RNeasy Midi kit (Qiagen, Valencia, CA, USA)
as per the manufacturer's instructions for animal tissues. In brief, 0.91 g of
embryos were homogenized in 12 ml of chaotropic buffer, and RNA was isolated
via selective binding to a silicagel-based membrane (Qiagen). For the
hour 24 samples, swimming nauplii were separated from the shed chorion using a
Buchner funnel. To prevent overloading of the membrane, only 0.76 g of nauplii
were used for each replicate. This reduced mass compensated for the loss of
the inert embryonic chorion during hatching. As an internal control, 1 µg
of synthetic RNA for strawberry chitinase was added to the chaotropic buffer
prior to each homogenization (see below for details). Homogenates were
centrifuged at 12 000 g for 10 min at 0°C to remove cell
debris, and 10 ml of the resultant supernatant was applied to the RNeasy spin
membrane. The same procedure was used to isolate RNA from diapause embryos.
Diapause mRNA used for RLM-RACE was isolated using this total RNA and an
Oligotex mRNA Mini kit (Qiagen).
The concentration of RNA in each sample was determined spectrophotometrically (A260). Purity of the RNA preparations was determined by spectrophotometric scan (250350 nm) of each sample. DNA contamination was determined by percent loss after treatment with Rnase-free DNase (RQ1; Promega) followed by acidic PCIA (phenol: chloroform: isoamyl alcohol) extraction and ethanol precipitation. Integrity of the RNA was examined by both northern blot analyses and ethidium bromide staining of 18 and 28 S ribosomal RNA bands on a denaturing agarose gel.
Production of synthetic RNA for northern controls and dot blot standards
A genomic DNA clone of chitinase II from strawberry
(Khan, 2002) was used for the
production of synthetic RNA to be used as an internal control for the A.
franciscana total RNA isolation. The clone consisted of 1002 base pairs
of genomic DNA ligated into a pGEM-T easy vector (Promega, Madison, WI, USA)
that had been digested with EcoRI. This plasmid construct was
linearized using PstI (Gibco-BRL, Gaithersburg, MD, USA) and
subjected to proteinase K digestion, phenol:CIA extraction and ethanol
precipitation. A MEGAscript in vitro transcription kit (Ambion) was
used to transcribe synthetic RNA with T7 polymerase. After treatment with
DNase I, the synthetic RNA was precipitated with lithium chloride and stored
as a precipitate at 80°C.
The same procedure was used to produce synthetic RNA from the A. franciscana V-ATPase B-subunit clone, originally amplified with degenerate primers. In this case, however, the plasmid construct was digested with SpeI (Gibco-BRL) prior to in vitro transcription, and the synthetic RNA was used as a positive control for the oligonucleotide probing of northern blots.
Northern and dot blots
Total RNA samples (5 µg) were dried in a Savant speedvac (Farmingdale,
NY, USA), heat denatured at 55°C for 15 min in 12 µl of denaturing
buffer (80% deionized formamide, 3.7% v/v formaldehyde and 1x MOPS
running buffer, pH 7.0), quick cooled in an ice slurry, and electrophoresed on
a 1.2% agarose gel containing 1x MOPS (pH 7.0) and 37% v/v formaldehyde.
Gels were rinsed in several washes of DEPC-treated water to remove
formaldehyde. RNA was transferred to a nylon membrane (Genescreen Plus;
Dupont, NEN, Boston, MA, USA) over a period of 17 h by capillary action using
10x SSPE buffer (0.1 mol l1
Na2HPO4/NaH2PO4 pH 7.0, 1.8 mol
l1 NaCl and 10 mmol l1 EDTA). After
transfer, membranes were rinsed with 5x SSPE, dried for 1 h at 60°C,
and the RNA was UV crosslinked to the membrane. Crosslinked blots were washed
in hybridization buffer (5x SSPE, 2% SDS, 10 µg ml1
Rnase-free calf thymus DNA and 50 µg ml1 yeast RNA),
pre-hybridized in the same buffer for 6 h at 65°C, and hybridized with a
32P-dCTP (NEN-PerkinElmer, Boston, MA, USA)-labeled cDNA probe (see
below) for 50 h at 65°C. The membranes were subsequently washed twice in
2x SSPE with 0.1% SDS, followed by a single wash in 0.2x SSPE with
0.1% SDS. The final blots were then wrapped in cellophane and exposed to a
phosphor screen. Imaging was conducted with either a STORM 840 phosphor imager
or a Typhoon 8600 variable mode imager from Molecular Dynamics (Sunnyvale, CA,
USA).
To quantify the strawberry synthetic RNA in A. franciscana samples
after isolation, 5 µg aliquot samples of total RNA were subjected to dot
blot analysis. Procedures followed a published protocol
(Hardewig et al., 1996) with
slight modification. All samples were combined with 85.8 µl of denaturing
reagent (80% deionized formamide, 3.7% formaldehyde and 2x SSPE, pH 7.0)
and 5 µg of yeast RNA, diluted to a final volume of 100 µl and heat
denatured at 55°C for 15 min. No loading dye was used. After the samples
were applied to the membrane, sample wells were rinsed with 300 µl of
2x SSPE. The membrane was then dried for 30 min at 70°C before UV
crosslinking. The hybridization procedure followed that outlined for northern
blots.
Northern and dot blot probes were labeled using a published protocol
(Hardewig et al., 1996). Probe
for B-subunit blots was produced using, as template, the 394 bp cDNA fragment
of the V-ATPase B-subunit originally amplified with degenerate primers. Probe
used for the strawberry control blots was produced using the strawberry
DNA/pGEM-T easy vector construct for template. The construct was subjected to
EcoR1 digestion prior to probe production and provided a template for
oligonucleotide probes capable of recognizing synthetic RNA synthesized from
the strawberry chitinase II clone.
Production of a recombinant protein for the V-ATPase B-subunit from A. franciscana
A region of the conserved central domain of the V-ATPase B-subunit from
Artemia franciscana, corresponding to base pairs 7281108, was
amplified for in-frame insertion into a pET 30a(+) expression vector (Novagen,
EMD Biosciences, San Diego, CA, USA) digested with EcoRI and
XhoI restriction enzymes. The insert was sequenced directly from the
plasmid construct to confirm in-frame orientation of the B-subunit fragment
insert with the T7 Lac Promoter and His Tag. The protocol for expression
followed instructions in the pET System Manual (10th edition; Novagen) for
pLysS host strain, BugBusters cell lysis, and denaturing inclusion body
isolation. Protocol for Ni-NTA column chromatography followed that outlined
for purification under denaturing conditions in the manual for His Bind kits
(Novagen). Column eluate was concentrated using a Centricon centrifugal filter
devise from Millipore (Billerica, MA, USA). Protein concentration was
determined spectrophotometrically (A280).
Western blots
A. franciscana embryos were sampled after 0, 4 and 8 h of
incubation at 2223°C. Pre-emergence embryos were dechorionated
immediately prior to homogenization, as previously described
(Kwast and Hand, 1993).
Manduca sexta midgut was removed and prepared for homogenization
according to Harvey et al.
(1990
). Embryos were weighed
and subjected to gentle homogenization in four volumes of buffer (300 mmol
l1 sucrose, 2 mmol l1 EDTA, 50 mmol
l1 Hepes, pH 7.5) using 59 passes with a
Teflon/smooth glass homogenizer from Thomas Scientific (Swedesboro, NJ, USA)
and Glas-Col motor (Terre Haute, IN, USA) set at a maximum of 2000 r.p.m.
Homogenates were subjected to differential centrifugation at 4°C to
isolate subcellular fractions for electrophoresis (crude supernatant, 10 min
at 1000 g; mitochondrial pellet, 15 min at 9000
g; heavy membrane pellet, 30 min at 48 000 g;
heavy microsomal vesicle pellet, 1 h at 100 000 g; microsomal
vesicle pellet and cytosolic supernatant, 1 h at 140 000 g).
Centrifugal forces were calculated for the maximum radius of JA 25.50 and Type
70.1Ti rotors (Beckman, Fullerton, CA, USA) used for centrifugation at
48
000 g and
100 000 g, respectively. All
centrifugation steps were performed in series using the supernatant of the
previous spin, and samples were kept at 4°C throughout the procedure.
Prior to western blotting, heavy and microsomal membrane preparations were
subjected to osmotic shock by resuspending pellets in 35 ml and 8 ml,
respectively, of a dilute buffer (10 mmol Tris-MOPS, pH 7.6). This procedure
served to remove trapped soluble protein from vesicles and helped verify that
dissociable V1 subunits visualized in western blots were indeed
associated with the membrane-spanning V0 domain prior to
electrophoresis. A second repetition of this procedure also served to reduce
the non-specific background observed in western blots hybridized with the
d-subunit antibody (Fig. 5C;
lane W).
|
Hatching studies
Prior to incubation in the treatment medium, embryos were rinsed briefly
with 35 artificial seawater at 22°C. Unless otherwise stated, all
embryos remained fully hydrated prior to incubation, and incubations were
performed in a 9:1 mixture of artificial sea water (35
) and 100%
ethanol. All hatching incubations were conducted in 12-well plastic cell
culture plates with 1025 embryos per well. A single plate was used for
each treatment, thus providing a total of 120300 embryos per treatment
group. In order to provide a high surface-to-volume ratio for gas exchange,
only 0.5 ml of the incubation medium was placed into each well prior to the
addition of embryos. To prevent evaporation of the treatment medium during
lengthy incubations, a wet paper towel was placed over the top of each plate
so that two ends of the towel were immersed in a distilled water bath. A
cardboard box was then placed over the entire setup to limit photooxidation of
the macrolide antibiotics used. Embryos were either allowed to develop
undisturbed for 6669 h before staging or were observed every 2 h over a
36 h period. All incubations were performed at 22°C.
Oligomycin (F-type ATPase inhibitor) was obtained from Sigma. The V-type
ATPase inhibitor bafilomycin A1 (also referred to as bafilomycin)
was either obtained from AG Scientific (San Diego, CA, USA) or LC Laboratories
(Woburn, MA, USA). Both oligomycin and bafilomycin were dissolved in 100%
ethanol, and the stock solutions were stored at 20°C. The
experimental concentrations of oligomycin and bafilomycin were varied and
their effect on hatching success determined. Under the final conditions used
(35 seawater + 10% ethanol), the limit of bafilomycin solubility was
observed to be 6 µmol l1. Because bafilomycin is unstable
in aqueous solution, the equivalent of 10 µmol l1
bafilomycin was used to ensure a maximal effect during prolonged
incubation.
Statistical analyses
All values are reported as means ± 1
S.E.M. Individual data points are given when
N<3. Unless otherwise noted, means for mRNA abundance were
compared among developmental time points using analysis of variance (ANOVA),
while post-hoc multiple comparisons were made using Tukey's
studentized range (HSD) test. Inclusion of hour 0 values in comparisons among
northern blot data summarized in Fig.
3B resulted in the violation of the assumption of homogeneity of
variance, as determined by Levene's test for homogeneity (F=6.71,
P=0.0033). Thus, comparisons between hour 0 and other time points
were performed separately using the KruskalWallis test, followed by the
Nemenyi test (Zar, 1999).
Statistical significance was set at P
0.05 for all analyses.
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Results |
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The putative start codon for the B-subunit of A. franciscana
embryos is preceded by a 70-nucleotide 5' untranslated region. The
initiation sequence surrounding the start codon (AAAATGA) resembles that of a
homologous subunit from the decapod crustacean Carcinus maenas
(GenBank accession no., AAF08281
Weihrauch et al., 2001) but
with an adenine at the #2 position instead of a cytidine. However, the A.
franciscana initiation sequence differs greatly from that of the optimal
Kozak initiation sequence (Kozak,
1991
), with a complete absence of cytosine immediately upstream of
the start codon and an adenosine in the #4 position. The stop codon begins at
nucleotide 1565 and is followed by a relatively short 216-nucleotide 3'
untranslated region. Only a single adenosine and uridine-rich element (ARE) is
present in this region, and polyadenylation begins at nucleotide 1772. The
exact length of the poly-A tail is unknown, due to the use of a poly-T primer
for reverse transcription. The complete transcript contains a single ORF,
encoding 498 amino acids. The encoded protein has a calculated pI of 5.28, and
its deduced molecular mass (55.5 kDa) closely matches values reported for
B-subunit isoforms from other species
(Filippova et al., 1998
;
Weihrauch et al., 2001
).
Definitive identification of the encoded protein was obtained via a
BLAST search of the Conserved Domain Database (CCD;
Marchler-Bauer et al., 2003
),
which revealed the presence of a highly conserved V-ATPase B-subunit NtpB
domain, spanning amino acid positions 26490
(Fig. 1).
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|
A second mRNA expression profile was produced in order to more rigorously
test for differential expression inferred from the trends above for the 2250
bp transcript (see Materials and methods). In this experiment, synthetic plant
RNA was used as an internal control for the examination of mRNA expression for
the V-ATPase B-subunit. The profile produced with this method clearly
demonstrates significant (H4=14.942) differential
expression of the 2250 bp band transcript during the first 24 h of development
(Fig. 3B). This B-subunit
message drops by 47.0±2.8% during the first 12 h of aerobic development
(Fig. 3B), at which point the
embryo is in the process of emerging from its protective cyst wall (emergence
stage 1; Fig. 3C). By 16 h of
development, message levels have decreased by 59.4±3.3% relative to
hour 0 (Fig. 3B), and the
embryo is now enclosed only by a thin hatching membrane (emergence stage 2;
Fig. 3C). By 24 h of
development, the swimming nauplius predominates
(Fig. 3C), and mRNA levels are
no longer significantly different from 0 h. Thus, our second experiment
confirms, with statistical significance, the trends seen for the 2250 bp
transcript in the first experiment. Importantly, this confirmation comes with
the use of two different RNA isolation procedures. The larger (3500 bp)
mRNA is also apparent in the northern blot of total RNA isolated by the RNeasy
method but was too faint to allow confirmation of the trends seen in its
expression from the first experiment (data not shown). It is noteworthy that a
single B-subunit gene can produce multiple mRNA transcripts with differing
sizes and developmental expression patterns. Work on the mosquito Culex
quinquefasciatus shows the expression of two B-subunit transcripts in
larvae (4.2 kb and 1.8 kb) but only one 3.0 kb transcript in pupae
(Filippova et al., 1998
). The
northern blot results presented here suggest that a similar pattern may exist
for A. franciscana. However, we were unable to identify more than one
distinct cDNA in these embryos despite the sequencing of multiple clones.
Analysis of V-ATPase protein expression during development
To determine whether or not V-ATPase protein is expressed during early
development, western blot analysis was performed on subcellular fractions
isolated during the first 8 h of development. Unlike that reported for the
decapod crustacean C. maenas
(Weihrauch et al., 2001),
A. franciscana blots hybridized with a monoclonal antibody raised
against the B-subunit from yeast (Molecular Probes, Eugene, OR, USA) produced
inconclusive results (data not shown). However, a polyclonal antibody raised
against the native V1 domain of the V-ATPase from M. sexta
(Huss, 2001
) did cross-react
with four proteins present in heavy membrane, microsomal vesicle and soluble
cytosolic fractions of A. franciscana lysates (Figs
4,
5A,B). These protein bands were
identified as subunits A, B, E and G of the V-ATPase by comparison with
immunoreactive proteins of similar size in lysates of M. sexta (Figs
4,
5B).
|
Identification of the 70 kDa A-subunit was confounded by the co-migration of a protein that gave a strong signal in all blots incubated with a biotinylated secondary antibody (Figs 4, 5A). For this reason, subunit A was visualized with a less-sensitive technique employing an alkaline phosphatase-conjugated secondary antibody (Fig. 5B). In these blots, expression of subunit A was detected in the microsomal fraction only (Fig. 5B, lane V140). In blots incubated with a polyclonal antibody recognizing the d-subunit from M. sexta, three immunoreactive bands are visible (Fig. 5C,D). While difficult to discern from nonspecific background fluorescence, these bands appear only on blots incubated with the primary antibody recognizing M. sexta d-subunit and never on control blots. Two of these bands are present in heavy membrane preparations only (Fig. 5C; 44.6 and 43.6 kDa bands), while the third is present in both heavy membrane and microsomal vesicle preparations (Fig. 5C; 40.8 kDa band). A comparison with the M. sexta homologue suggests that the third, or smallest, of these bands is the d-subunit from the V-ATPase of A. franciscana (Fig. 5D). This band does not appear in the membrane-free cytosolic fraction.
Three immunoreactive bands (>105, 94 and 60 kDa) are also visible in blots of Artemia lysates incubated with a polyclonal antibody recognizing the a-subunit from M. sexta (Fig. 5E). The 94 kDa band was identified as subunit a based on size comparison with the M. sexta homologue. The >105 kDa band may be an aggregate with a second protein, as heat denaturing the sample for >1 min causes the 94 kDa band to disappear while the >105 kDa band increases in intensity for both M. sexta and A. franciscana samples (data not shown). The 60 kDa band is most likely to be a breakdown product of the 94 kDa a-subunit. All three of these bands appear in western blots of M. sexta membrane preparations (Fig. 5E, lane Ms).
Abundance of the membrane-bound d- and a-subunits from the V0 domain does not change appreciably during the first 8 h of development (Fig. 5E). This finding stands in contrast to that for three of the soluble subunits of the V1 domain (B, E and G), which show a small qualitative increase in expression within the heavy membrane fraction (Fig. 5A, lane H), but not in the larger cytosolic pool (Fig. 5A, lane S140), over the same time period.
Effect of V-ATPase and ATP-synthase inhibitors on hatching success
Prior to inhibitor studies, tests were conducted to determine whether or
not the ethanol used to solubilize inhibitors impacted hatching of embryos in
varying salinities of artificial seawater. Ethanol had no effect on the
hatchability of dechorionated embryos at concentrations of 10%
(Fig. 6). Concentrations in
excess of 10% eventually caused bleaching of color and mortality, which
indicates permeability of embryos to ethanol. It is of interest to note that
the effect of ethanol and macrolide antibiotics on hatching success of
dechorionated embryos was affected by the severity of the dechorionation
procedure. Both the age of the dechorionation solution and variation in the
exposure to this solution influenced survivorship of subsequent treatments
even though differences in embryos were not detected with light or scanning
electron microscopy (J.A.C. and S.C.H., unpublished observations).
|
The permeability of encysted A. franciscana embryos to lipid-soluble inhibitors was determined by observing hatching success in varying concentrations of oligomycin, a macrolide antibiotic that specifically inhibits the F-type ATP-synthase of mitochondria. The ability of dechorionated embryos to develop beyond the encysted stage decreased as the concentration of oligomycin was increased. In contrast to the results presented below for bafilomycin, only an extremely small number of individuals developing beyond this stage ceased to develop during the subsequent E1 or E2 stages of emergence (Fig. 7A). Similar results were obtained if the dechorionated embryos were blotted dry prior to incubation, but the decrease in hatching success was more pronounced at lower concentrations of oligomycin (Fig. 7B), probably due to greater inhibitor uptake resulting from bulk flow of water during rehydration. By contrast, hatching of control embryos with an intact chorion was unaffected by oligomycin concentrations as high as 10 µmol l1 (Fig. 7B).
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Discussion |
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In order to better appreciate the role that acidic compartments might play
in anoxia signaling, we examined the potential locations of such compartments
within the anoxia-tolerant embryos by focusing on expression of the V-ATPase
proton pump. All attempts to localize the V-ATPase using immunohistochemistry
were unsuccessful. Thus, we employed a combination of sequence data analysis
and western blotting to infer its subcellular distribution. Sequence
alignments (Fig. 1) and
phylogenetic analysis (Fig. 2)
demonstrate that cDNA for the V-ATPase B-subunit of A. franciscana
(Crustacea; Anostraca) groups with homologous B-subunit isoforms from
invertebrates specifically between the decapod crustacean Carcinus
maenas and insects. A breakdown of this phylogenetic analysis produces
two distinct groupings: `generalist' and `specific', based on the subcellular
localization and functional repertoire of the holoenzyme. The `specific'
grouping includes mammalian B-subunit isoforms known to be incorporated into
V-ATPase employed specifically for proton secretion across the plasma
membrane. By contrast, the `generalist' grouping includes invertebrate and
mammalian isoforms likely to be incorporated into V-ATPase present in both
intracellular and plasma membranes for a diverse array of functions
(Futai et al., 2000). Given
that the A. franciscana isoform groups within the generalist clade,
it is plausible that the V-ATPase of the anoxia-tolerant embryos is localized
to both plasma and organellar membranes. Indeed, the western blot data
presented here bolster this hypothesis, supporting the argument for a broadly
localized distribution of V-ATPase within encysted embryos. The
co-localization of V1 domain subunits (B, E and G) with
V0 domain subunits (d and a) in both heavy and microsomal membrane
preparations (Fig. 5) is
consistent with the distribution of intact V-ATPase in both plasma and
vesicular membranes during the earliest stages of encysted development.
However, it should be noted that V-ATPase localization to the plasma membrane
can be the result of participation in exocytosis or membrane energization,
neither of which involve the establishment of a large proton chemical gradient
(Harvey and Wieczorek, 1997
).
Thus V-ATPase-acidified compartments are likely to be intracellular.
An analysis of the V-ATPase expression data presented here in the context
of recent literature strongly suggests a role for this enzyme in the
establishment of intracellular proton stores in developing embryos. Wieczorek
et al. (2000) noted that
biosynthesis of V-ATPase subunits is tightly regulated in instances when the
V-ATPase functions in a capacity other than the acidification of intracellular
compartments. These authors point out that a molting-induced decrease in the
activity of V-ATPase from Manduca sexta midgut parallels decreases in
transcript levels for every subunit examined. However, for at least the first
8 h of development, such parallel changes are not apparent in A.
franciscana, as the abundance of B-subunit mRNA during pre-emergence
development (Fig. 3B) is not
tightly linked to the level of the encoded protein incorporated into the
holoenzyme (Fig. 5). Abundance
of the 2250 bp B-subunit mRNA decreases consistently throughout the first 16 h
of development (Fig. 3B), while
western analyses suggest a modest increase in the incorporation of B-subunit
protein in membrane-bound V-ATPase complexes. Based on these observations, one
might suggest that acidification of intracellular compartments is the primary
role for the V-ATPase of brine shrimp embryos. Indeed, published data
regarding the development of A. franciscana embryos offer some
support for this hypothesis. Increases in enzyme activities associated with
transcription (Slegers, 1991
)
and in total protein labeling from pulsechase experiments
(Peterson et al., 1978
)
suggest an elevated demand for trafficking of newly synthesized proteins prior
to emergence of the brine shrimp larva. In addition, both an embryonic cuticle
and the first naupliar exoskeleton are synthesized by the epithelial cells
during the process of emergence (Rosowski
et al., 1997
), suggesting the need for a very active exocytotic
pathway between 8 and 24 h of development. Lysosomal activity has also been
noted to be involved in the degradation of yolk platelets
(Perona et al., 1988
;
Perona and Vallejo, 1989
).
Given that protein trafficking, yolk platelet degradation and exocytosis
require V-ATPase-mediated acidification of lysosomes, transport vesicles and
Golgi, it seems probable that the acidification of these intracellular
compartments is a primary function of the brine shrimp enzyme during early
development.
If proton chemical gradients are involved in anoxia signaling in brine
shrimp embryos, a mechanism would be required to coordinate oxygen sensing
with net proton flux from acidic compartments. Recently published work on
vertebrate, insect and yeast V-ATPase suggests that the V-ATPase of brine
shrimp embryos may be downregulated under anoxia by changes in ATP:ADP ratio
and inorganic phosphate (Pi) concentration. For this reason, it is
of interest that A. franciscana embryos deal with repeated and
prolonged bouts of environmental anoxia by entering a reversible state of
metabolic depression characterized, in part, by a reverse Pasteur effect.
Under anoxia, these embryos rapidly shut down both catabolic and anabolic
processes (Hand and Hardewig,
1996) and ultimately approach an ametabolic state
(Warner and Clegg, 2001
).
Concomitant with this metabolic downregulation is a precipitous drop in
adenylate status (Stocco et al.,
1972
). Over a 24 h period of anoxia, ATP levels decrease by
94.6±0.9% (Anchordoguy and Hand,
1994
), with the majority of this decline occurring in the first
few minutes. As a result, during the first 21 min of anoxia, the ATP:ADP ratio
decreases from 7.6 to 0.92 (Anchordoguy and
Hand, 1994
) while [Pi] increases greatly
(Rees et al., 1989
).
Importantly, recent catalytic analysis of V-ATPase from both insects and yeast
demonstrates that even small reductions in the ATP:ADP ratio inactivate this
proton pump (Huss and Wieczorek,
2003
; Kettner et al.,
2003
), while work on vertebrates and yeast shows that both ADP
(Kettner et al., 2003
;
Vasilyeva and Forgac, 1998
)
and Pi (David and Baron,
1994
) have inhibitory effects on V-ATPase activity as well. If
regulation by such biochemical variables occurs in A. franciscana,
proton translocation by the V-ATPase would stop within minutes of anoxic
exposure. Provided that all H+ gradient dissipative paths are not
concurrently downregulated, directional net H+ flux would then
dissipate existing gradients and acidify the surrounding cytoplasm.
Interestingly, incubation of permeabilized mammalian cells in ATP-free medium
resulted in a 0.7 pH unit alkalinization of the trans-Golgi network, which was
fully reversed by the addition of exogenous ATP
(Demaurex et al., 1998
). These
authors also demonstrate an alkalinization of Golgi during incubation with the
V-ATPase inhibitor conconamycin. Similar effects were observed with
bafilomycin incubation or ATP deprivation for yolk platelets of permeabilized
Xenopus oocytes (Fagotto and
Maxfield, 1994b
). Taken together, these data suggest that
inhibition of V-ATPase activity via a decrease in cellular ATP is
enough to cause a net proton flux from organelles such as Golgi and yolk
platelets to the cytosol.
Leakage of protons from intracellular compartments can significantly
acidify the cytosolic space (Madshus et
al., 1987). However, in any given cell, the effect of proton flux
from these compartments would ultimately depend on their relative buffering
capacities and size in relation to the surrounding cytoplasmic space. While we
are unaware of any studies reporting these values in Artemia embryos,
a large body of literature does describe such values for components of the
protein synthetic, degradative, exocytotic and endocytotic pathways of other
species (Ibarrola et al.,
2000
; Kelly et al.,
1991
; Rybak et al.,
1997
; Schoonderwoert and
Martens, 2001
; Van Dyke and
Belcher, 1994
; Wu et al.,
2001
). Based both on these data and published morphological
studies on Artemia (Perona et
al., 1988
; Perona and Vallejo,
1989
), one can estimate that Golgi, lysosomes and various
transport vesicles constitute 2% of the intracellular volume in
Artemia embryos and have a weighted average pH of 5.4 with a
buffering capacity of 60 mmol H+ l cytosol1 pH
unit1 over the predicted range of compartment alkalinization
under anoxia. It is also reasonable to include an estimate of the effect of
proton release from membrane-bound yolk platelets in Artemia, the
potential acidification of which has been discussed elsewhere
(Busa et al., 1982
;
Fagotto, 1991
). Because the
yolk platelets of Artemia are most stable in vitro at a pH
of 5.7 (Utterback and Hand,
1987
), and an in vivo pH of 5.6 was observed for
non-degrading yolk platelets in Xenopus oocytes
(Fagotto and Maxfield, 1994a
),
it is plausible to assume a pH of 5.7 for the yolk platelets of A.
franciscana embryos. We can also conservatively assume a buffering
capacity of 60 mmol H+ l cytosol1 pH
unit1 given that 80% of embryonic protein is stored in the
yolk platelets (Vallejo et al.,
1981
) and that yolk platelets of other species are known to have a
high buffering capacity (Fagotto and
Maxfield, 1994b
). A two-dimensional analysis of electron
micrographs generated by our lab (data not shown) suggests that approximately
15% of the embryonic space is occupied by yolk platelets. Using these data in
conjunction with a general cytoplasmic buffering capacity of 18 mmol
H+ l cytosol1 pH unit1
published by Kwast et al.
(1995
), and assuming that
intracellular compartments will not alkalinize above pH 6.4 under anoxia, one
can estimate the contribution of protons from these compartments to be 50% of
that required to explain the pH shift occurring during the first 20 min of
anoxia
.
In summary, it seems improbable that a concentration gradient of protons
could be maintained across lipid bilayers over years of anoxia in the face of
an absence of available cellular energy for proton transport. We propose that,
rather than undergoing a slow gradient dissipation spanning months or years,
A. franciscana embryos experience an acute release of protons from
acidic compartments, which is triggered by exposure to anoxia. Inactivation of
H+ pumping by the V-ATPase would be an essential step for such a
process and may represent an adaptive trait critical to anoxia tolerance in
this species. The data reported here demonstrate that this proton pump is
differentially expressed during the period of encysted development
characterized by anoxia tolerance, while inhibition of its activity with
bafilomycin halts embryonic development. Together, these data demonstrate a
critical need for proton pumping prior to the rapid growth phase accompanying
emergence. Interestingly, the sharp decline in ATP:ADP ratio seen in A.
franciscana under anoxia could inactivate the V-ATPase within minutes of
exposure to anoxia. Subsequent dissipation of proton chemical gradients could
help facilitate the observed cytoplasmic acidification required for
quiescence. In the accompanying paper
(Covi et al., 2005), we
present 31P-NMR experiments that directly test the ability of
proton gradient dissipation to acidify the intracellular space.
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