799
Depression of Protein Synthesis during Diapause in Embryos of the
Annual Killifish Austrofundulus limnaeus
Jason E. Podrabsky*
Steven C. Hand†
Section of Integrative Physiology and Neurobiology,
Department of Environmental, Population, and Organismic
Biology, University of Colorado, Boulder, Colorado 803090334
Accepted 7/12/00
ABSTRACT
Rates of protein synthesis are substantially depressed in diapause II embryos of Austrofundulus limnaeus. Inhibition of oxygen consumption and heat dissipation with cycloheximide indicates that 36% of the adenosine triphosphate (ATP) turnover
in prediapausing embryos (8 d postfertilization [dpf]) is caused
by protein synthesis; the contribution of protein synthesis to
ATP turnover in diapause II embryos is negligible. In agreement
with the metabolic data, incorporation of amino acids (radiolabeled via 14CO2) into perchloric acid–precipitable protein decreases by over 93% in diapause II embryos compared with
embryos at 8 dpf. This result represents a 36% reduction in
energy demand because of depression of protein synthesis during diapause. Adjusting for changes in the specific radioactivity
of the free amino acid pool at the whole-embryo level yields
rates of protein synthesis that are artifactually high and not
supportable by the observed rates of oxygen consumption and
heat dissipation during diapause. This result indicates a regionalized distribution of labeled amino acids likely dictated
by a pattern of anterior to posterior cell cycle arrest. AMP/ATP
ratios are strongly correlated with the decrease in rates of protein synthesis, which suggests a role for adenosine monophosphate (AMP) in the control of anabolic processes. The major
depression of protein synthesis during diapause II affords a
considerable reduction in energy demand and extends the duration of dormancy attainable in these embryos.
* To whom correspondence should be addressed. Present address: Hopkins Ma-
rine Station, Stanford University, Oceanview Boulevard, Pacific Grove, California
93950-3094; e-mail: [email protected].
†
Present address: Department of Biological Sciences, Louisiana State University,
Baton Rouge, Louisiana 70803.
Physiological and Biochemical Zoology 73(6):799–808. 2000. q 2000 by The
University of Chicago. All rights reserved. 1522-2152/2000/7306-99122$03.00
Introduction
Populations of the annual killifish, Austrofundulus limnaeus,
survive seasonal drying of their ephemeral pond habitats by
producing drought-tolerant, diapausing embryos (Wourms
1972a, 1972b). Diapause is a state of developmental arrest that
precedes the onset of unfavorable environmental conditions
(Hand 1991; Hand and Podrabsky 2000). Diapause is promoted
by endogenous cues and typically occurs as part of the natural
developmental program; thus, embryos may enter diapause
even under conditions considered optimal for development.
Diapausing embryos of A. limnaeus may remain dormant for
extended periods of time, spanning periods from months to
years (Wourms 1972b; Podrabsky 1999). While there may be
three distinct stages of diapause in some annual killifish
(Wourms 1972b), diapause interrupts development in embryos
of A. limnaeus at only two distinct developmental stages (diapause II and III; Podrabsky 1999; Podrabsky and Hand 1999).
At 257C diapause II occurs at 24 d postfertilization (dpf) in an
embryo possessing 38–40 pairs of somites, the foundations of
the central nervous system, optic cups, otic vesicles, and a
functional tubular heart (Wourms 1972a; Podrabsky 1999).
Diapause III occurs in the fully developed embryos just before
hatching and has been compared with estivation in African
lungfish (Wourms 1972b). Of the two diapause stages that occur
in A. limnaeus, diapause II exhibits the more profound depression of metabolism (Podrabsky and Hand 1999) and appears to be more resistant to environmental stresses such as
anoxia and dehydration (J. E. Podrabsky, unpublished data).
A 90% depression of metabolism accompanies diapause II
(compared with embryos at 8 dpf) in embryos of A. limnaeus,
and a simultaneous downregulation of ATP-producing and
ATP-consuming processes is indicated by the maintenance of
stable levels of adenosine triphosphate (ATP) during the transition in metabolic rate (Podrabsky and Hand 1999). One prediction from these findings is that major biosynthetic events
are downregulated during diapause. In this study, we tested the
hypothesis that decreased rates of protein synthesis may reduce
energy demand and thus facilitate the metabolic depression
observed in diapause II embryos of the annual killifish A.
limnaeus.
It should be noted that developmental arrest is not necessarily
associated with a decrease in metabolic rate in all cases. For
800 J. E. Podrabsky and S. C. Hand
example, embryos of the California grunion, Leuresthes tenuis
(Darken et al. 1998), and some amphibian embryos (Bradford
and Seymour 1985) enter a state termed “delayed hatching”
that is not coupled to a depression in metabolic rate. Similarly,
when embryos of the brine shrimp Artemia franciscana enter
diapause (as judged by arrest of cell division and morphogenesis), metabolism is high and comparable to active cysts (Clegg
et al. 1996). Thereafter, the rates of oxygen consumption and
protein synthesis decrease slowly over several days to low values
(Clegg et al. 1996). Thus, cessation of development in embryos
of A. franciscana is not temporally linked with metabolic
downregulation.
When metabolic depression is observed, a coordinated arrest
of both catabolic and anabolic processes appears to be a common trait of organisms that can enter and recover from such
states (Hochachka and Guppy 1987; Storey and Storey 1990;
Guppy et al. 1994; Hand and Hardewig 1996; Hand 1999).
Protein biosynthesis is one of the most energetically expensive
processes in the cell, and thus, a reduction in the rate of protein
synthesis may be essential in the face of severe metabolic depression if energy balance is to be maintained. Estimates indicate that protein synthesis may account for 12%–40% of the
basal metabolism of mammalian cells, tissues, or whole animals
(Buttgereit et al. 1992; Buttgereit and Brand 1995; Rolfe and
Brown 1997; Fuery et al. 1998a). Protein synthesis is estimated
to account for as high as 60%–90% of the total oxygen consumption of various cell lines and primary cell cultures isolated
from fish (Pannevis and Houlihan 1992; Smith and Houlihan
1995). The reduction in energy demand resulting from the
depression of protein synthesis during dormancy can be substantial. Fuery et al. (1998b) estimate that 52% of the metabolic
depression observed in liver tissue isolated from the estivating
frog Neobatrachus centralis is caused by a decrease in the rate
of protein synthesis. Land et al. (1993) report a 92% decrease
in the rate of protein synthesis of isolated turtle hepatocytes
when exposed to anoxia, and this arrest accounts for nearly
36% of the metabolic depression observed.
Our results indicate a global arrest of protein synthesis. While
this arrest is a major mechanism of energy conservation during
diapause II and accounts for nearly 36% of the overall metabolic
depression, it may restrict the embryo’s ability to respond to
changes in the internal milieu or external environment via de
novo gene expression. This limitation may have a profound
influence on the nature and timing of mechanisms that sustain
diapause and afford embryos tolerance to environmental
perturbations.
Material and Methods
Developmental Stages Investigated
On the basis of previous data (Podrabsky and Hand 1999), five
embryonic stages were chosen for investigation in this study:
2, 8, 16, 24, and 42 dpf. These stages span the major metabolic
states associated with early development and diapause II. Embryos at 2 dpf (late blastula, early epiboly) exhibit an increasing
metabolic rate. Metabolism peaks during dispersion and reaggregation at 8 dpf, about 2 d before the formation of the embryonic axis. The metabolism of embryos at 16 dpf is about
one-half the metabolism of embryos at 8 dpf. Embryos at 16
dpf have a functional heart and half the number of somites
present in diapause II embryos. Development ceases when embryos enter diapause II at 24 dpf, and at this point, metabolism
is depressed 67% relative to embryos at 8 dpf. Oxygen consumption continues to decline during diapause II, and by 42
dpf, it is depressed by 90% compared with 8 dpf.
Measurements of Protein Synthesis
Embryos of Austrofundulus limnaeus are virtually impermeable
to free amino acids (J. E. Podrabsky, unpublished data). Thus,
the free amino acid pool was radiolabeled by heterotrophic
fixation of 14CO2 (cf. Hofmann and Hand 1990). Heterotrophic
fixation of CO2 in animal cells is caused by a number of enzymes
that produce intermediates of the citric acid cycle, including
phosphoenolpyruvate carboxykinase (Lowenstein 1967). Radioactivity fixed in this manner is recovered largely (∼80%) as
amino acids because of transamination (Clegg 1976). Typically,
as in our study, 185% of the radioactivity present in the free
amino acid pool is found in aspartic acid and glutamic acid
(Hofmann and Hand 1990). Determination of the rate of protein synthesis is based on measuring the incorporation of these
in vivo radiolabeled amino acids into acid-precipitable protein.
Embryos were collected and incubated as previously described (Podrabsky 1999) before use in radiolabeling experiments. On the day of experimentation, the embryos were aseptically cleaned with a solution of 0.01% sodium hypochlorite
as described in Podrabsky (1999). Embryos were divided into
groups of 30 and transferred to a 2-mL microcentrifuge tube
that contained 1 mL preincubation solution, which consisted
of embryo medium (Podrabsky 1999) fortified with 25 mM
HEPES buffer (pH 7.4) and 20 mg L21 gentamycin sulfate with
or without 10 mM cycloheximide. Embryos were preincubated
for 20 min in this solution at 257C. The assay was started by
replacing the preincubation solution with 1 mL of labeling
medium composed of preincubation medium plus 50 mCi mL21
of H14CO3 (Dupont/NEN, Boston). Cycloheximide (10 mM)
was added to the appropriate tubes as in the preincubation.
The microcentrifuge tubes were closed and incubated at 257C
for 1–4 h. Tubes were inverted every 10–15 min to mix the
labeling medium. At selected time points, embryos were removed, blotted briefly on a paper towel, and homogenized in
300 mL of ice-cold deionized water in a ground glass tissue
homogenizer. Triplicate aliquots (30 mL each) of the homogenate were applied to Whatman GF/C glass fiber filters. The
remaining homogenate was then stored at 2807C for later determination of specific radioactivity in free amino acids (see
Depression of Protein Synthesis during Diapause 801
Table 1: Concentration-dependent inhibition
of amino acid incorporation into protein by
cycloheximide in 8-d postfertilization
embryos
Cycloheximide (mM)
Percentage Inhibition
20 .......................
10 .......................
5 .........................
75.2
79.1 5 .3A
72.0 5 .4B
Note. Values are means 5 SEM (n p 2 for 20 mM;
n p 4 for 5 and 10 mM). Values were calculated from total
incorporation after 4 h of incubation. Means with the same
letter are not statistically different (Tukey’s HSD, P ! 0.05).
below). The filters (90–120) were then plunged into 500 mL
of 10% trichloroacetic acid (TCA) with 5 mM each of aspartic
acid and glutamic acid. After 10 min, the filters were placed in
500 mL of 5% TCA at room temperature for 15 min. After a
second wash in 5% TCA as above, the filters were transferred
to 250 mL of 95% ethanol for 10 min at room temperature.
The filters were allowed to air dry for 1–3 h. Blank filters were
always processed with the filters containing radiolabeled proteins to correct for background binding of radioactivity to the
filters. Each filter was added to a scintillation vial with 2 mL
of Scintiverse II. The amount of radioactivity on each filter was
determined using a liquid scintillation counter (LKB/Wallac,
Gaithersburg, Md.). Counting efficiencies were uniform across
all samples (data not shown), and thus, counts per minute
(CPM) is directly proportional to disintegrations per minute
(DPM) for all samples. Parallel values determined in the presence of cycloheximide were subtracted from each experimental
sample.
Specific Radioactivity of Free Amino Acids
The specific radioactivity of the free amino acid pool was determined for each sample. Embryo homogenates (previously frozen) were allowed to thaw on ice. Perchloric acid (PCA) extracts
were prepared from the homogenate by the addition of 70%
PCA (final concentration of 1 N PCA) to precipitate proteins.
The acid homogenate was then centrifuged at 10,000 g for 10
min at 47C to remove the protein. The supernatant was neutralized with 5 M K2CO3. The neutralized supernatant was centrifuged as above to remove the perchlorate salts, and 40 mL of
the homogenate were added to 80 mL of o-phthalaldehyde reagent
(Fluoraldehyde solution, Pierce, Rockford, Ill.) to derivatize the
amino acids. The solution was mixed and incubated at room
temperature for 2 min. The reaction was stopped by the addition
of 80 mL of ice-cold 0.1 M sodium acetate (pH 7.0). The sample
was immediately filtered (0.45 mm), and 100 mL were analyzed
for amino acids.
Amino acids were quantified by reverse-phase, high-performance liquid chromatography (HPLC, Milton Roy 4000 series,
Rochester, N.Y.) as described in Kwast and Hand (1993). Derivatized samples were separated using an APEX I octadecyl column (250 # 4.6 mm, particle size 5 mm; Jones Chromatography,
Lakewood, Colo.). Separation of amino acids was achieved using
a mobile phase gradient. Solution A consisted of 0.1 M sodium
acetate, pH p 7.7 (titrated with 1 N HCl), methanol, and tetrahydrofuran in a ratio of 90 : 9.5 : 0.5 (v/v). Solution B was
100% methanol. The solvent gradient was adjusted for optimal
separation of each amino acid as follows (expressed as the percentage of Solution A, with the balance being Solution B): 0–2
min, 95%–90% solvent A; 2–15 min, 90% A; 15–24 min,
90%–60% A; 24–35 min, 60%–40% A; 35–39 min, 40% A; 39–42
min, 40%–0% A; 42–46 min, 0% A; 46–48 min, 0%–95% A.
The flow rate of 1.5 mL min21 was constant throughout the
gradient. Amino acid peaks were detected at 330 nm and were
quantified by integration of peak area. Fractions (0.75 mL volume) were collected for the first 10 min of each chromatographic
separation using a Rainin model FC-100 fraction collector. Aspartic acid and glutamic acid accounted for 195% of the radioactivity associated with free amino acids (data not shown). For
this reason, calculations of specific radioactivity were restricted
to data for aspartic acid and glutamic acid. Radioactivity in the
Table 2: Regression analysis of radiolabeled
amino acid incorporation during 4-h assays in
the presence and absence of 10 mM
cycloheximide
Age of Embryos (Days
Postfertilization)
and Incorporation
2 dpf:
Total ..................
Cycloheximide ......
8 dpf:
Total ..................
Cycloheximide ......
16 dpf:
Total ..................
Cycloheximide ......
24 dpf:
Total ..................
Cycloheximide ......
42 dpf:
Total ..................
Cycloheximide ......
Adjusted r 2
Slope
.85
.76
1,466A
98.9B
.97
.95
4,431A
1,280B
.92
.96
1,047A
349B
.95
.91
478A
182B
.91
.89
302A
129B
Note. All slopes (CPM mg21 DNA h21) are significantly different from zero (P ! 0.0001, Fisher’s PLSD). Slopes with the
same letter within each developmental stage are not significantly different based on a t-test comparing the slopes (P !
0.05) and the presence of a significant interaction term using
ANCOVA analysis with incubation time as the covariate
(P ! 0.0001).
802 J. E. Podrabsky and S. C. Hand
Table 3: Inhibition of incorporation of
radioactive amino acids into protein by 10
mM Cycloheximide as a function of
developmental stage
Days Postfertilization
Percentage Inhibition
2 ........................
8 ........................
16 .......................
24 .......................
42 .......................
95.3
77.9
86.9
84.4
85.2
5
5
5
5
5
1.2A
1.4B
1.0C
2.1B,C
2.0C
Note. Values for inhibition are means 5 SEM (n p 4).
Values were calculated from total incorporation after 4 h
of incubation. Means with the same letter are not statistically different (Tukey’s HSD, P ! 0.05).
appropriate fractions was determined using liquid scintillation
counting.
was measured for an additional 40–60 min in the presence of
inhibitor.
Statistics
All statistical analyses were performed using SAS 6.0 (SAS
Institute 1997) or StatView 5.0 (SAS Institute 1998) software.
All time-course data were evaluated via regression analysis.
Regression coefficients for incorporation of radiolabel in the
presence and absence of cycloheximide were compared using
a t-test and by testing for a significant interaction term in
ANCOVA analysis with time as the covariate. All other comparisons were made using ANOVA analysis. Post hoc comparisons were made with either Tukey’s honestly significant
difference (HSD) test or Fisher’s protected least significant
difference (PLSD) test.
Results
Amino Acid Incorporation Into Protein
Cycloheximide-Inhibitable Respiration and Heat Dissipation
Oxygen consumption was determined at 257C using polarographic oxygen electrodes (model 1302, Strathkelvin Instruments, Glasgow, Scotland) as previously described (Podrabsky
and Hand 1999). Seventy embryos were assayed in 1.5 mL of
air-saturated embryo medium containing 10 mg L21 gentamycin sulfate for 30–40 min. The embryo medium was continuously stirred during measurement of oxygen consumption
with a glass-coated magnetic stir bar. Embryos were held above
the stir bar using a fiberglass mesh platform. The chamber was
then opened, flushed with embryo medium containing 10 mg
L21 gentamycin sulfate and 10 mM cycloheximide, and closed
again for an additional 30–40 min to record cycloheximideinhibited oxygen consumption. Opening and closing the respirometry chamber had no effect on the oxygen consumption
of embryos (data not shown). The concentration of cycloheximide (10 mM) used in measurements of oxygen consumption
and heat dissipation was optimized for maximal inhibition of
protein synthesis based on radioactive incorporation experiments (see “Results”; Table 1).
Heat dissipation was measured at 257C as described in Podrabsky and Hand (1999). Briefly, 70 embryos were placed in a
stainless steel ampoule containing embryo medium supplemented as described above for determination of oxygen consumption. Embryo medium was equilibrated to a gas mixture
of 40% oxygen and 60% nitrogen. The elevated oxygen prevented embryos from experiencing hypoxia in the absence of
stirring. After the ampoule temperature equilibrated with the
measuring cylinder (typically 30 min), control rates of heat
dissipation were recorded for 40–60 min. The ampoule was
then removed, and the embryo medium was replaced with fresh
medium containing 10 mM cycloheximide. Heat dissipation
Embryos (8 dpf) were incubated in various concentrations of
cycloheximide to assess the efficacy of cycloheximide as an
inhibitor of protein synthesis in intact embryos. Maximal inhibition is observed with 10 mM cycloheximide (Table 1). For
all subsequent experiments, this level of cycloheximide was used
to inhibit protein synthesis. While this concentration is an order
of magnitude higher than is typically used to inhibit protein
synthesis in isolated cells (e.g., Buttgereit and Brand 1995), the
increased level required here is not surprising considering the
low permeability of these embryos to amino acids. Consequently, it is unlikely that the embryonic cells actually experienced the full dosage of cycloheximide.
During the 4-h assays, incorporation of radiolabeled amino
acids increases in a linear manner in both the presence and
Figure 1. Cycloheximide-inhibitable incorporation of radiolabeled
amino acids into protein. The slopes of all regressions are significantly
greater than 0 (P ! 0.0001). Filled symbols (mean 5 SEM, n p 4) represent developing stages, while open circles denote diapause II stage
embryos.
Depression of Protein Synthesis during Diapause 803
Table 4: Free amino acids in whole and yolk-free embryos of Austrofundulus limnaeus as a function of
development through diapause II
Whole Embryosa
Yolk-Free Embryosb
Amino Acid
2 dpf
8 dpf
16 dpf
24 dpf
42 dpf
8 dpf
16 dpf
24 dpf
42 dpf
Alanine ..............
.499
(.052)
.238
(.027)
.035
(.011)
.855
(.068)
.438
(.060)
.600
(.071)
.291
(.039)
.010
(.010)
.102
(.021)
.190
(.038)
.464
(.031)
.066
(.017)
.072
(.021)
.304
(.095)
ND
1.48
(.138)
1.48
(.064)
.577
(.010)
.352
(.032)
2.39
(.027)
17.7
(.306)
1.07
(.073)
.739
(.002)
.775
(.023)
.996
(.045)
3.51
(.067)
.262
(.030)
.591
(.037)
3.27
(.070)
.914
(.226)
.247
(.022)
.784
(.056)
1.15
(.030)
2.32
(.090)
1.74
(.022)
.736
(.028)
.278
(.019)
3.38
(.162)
20.4
(.129)
1.07
(.107)
.742
(.106)
.731
(.019)
.882
(.036)
4.36
(.153)
.243
(.039)
.763
(.046)
4.13
(.133)
.879
(.126)
.279
(.019)
.858
(.069)
1.14
(.020)
3.41
(.121)
1.92
(.091)
.717
(.025)
.212
(.010)
3.96
(.187)
21.2
(.562)
1.06
(.129)
1.06
(.076)
.502
(.020)
.388
(.027)
4.50
(.296)
.132
(.010)
.679
(.044)
4.15
(.301)
.714
(.166)
.232
(.017)
.642
(.088)
.846
(.053)
.016
(.001)
ND
.065
(.003)
ND
.071
(.006)
ND
.063
(.002)
ND
ND
ND
ND
ND
.011
(.001)
ND
.020
(.001)
ND
.024
(.002)
ND
.019
(.000)
ND
.075
(.006)
ND
.155
(.013)
ND
.130
(.015)
ND
.149
(.015)
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
.032
(.002)
ND
.047
(.002)
ND
.058
(.006)
ND
.052
(.003)
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
.035
(.020)
.079
(.003)
.243
(.049)
.720
(.051)
.937
(.052)
.580
(.076)
.381
(.016)
1.20
(.078)
10.7
(.365)
.397
(.140)
.344
(.066)
.669
(.029)
1.09
(.073)
1.84
(.272)
.178
(.060)
.238
(.033)
1.815
(.110)
.452
(.055)
.152
(.021)
.493
(.017)
.913
(.068)
ND
ND
ND
ND
.009
(.001)
ND
.020
(.007)
ND
.019
(.001)
ND
.024
(.006)
ND
4.51
(.470)
22.3
(1.40)
37.9
(1.04)
45.7
(.913)
46.7
(1.99)
NA
NA
NA
NA
Arginine .............
Asparagine ..........
Aspartic acid .......
Glutamine ..........
Glutamic acid ......
Glycine ..............
Histidine ............
Isoleucine ...........
Leucine ..............
Lysine ................
Methionine .........
Phenylalanine ......
Serine ................
Threonine ..........
Tryptophan .........
Tyrosine .............
Valine ................
Totals .............
Note. ND (not detectable) indicates values were below the detection limit of the instrument. NA (not applicable) indicates that total
free amino acids were not calculated for yolk-free embryos because of incomplete survey.
a
Values are nmol embryo21 (SEM in parentheses) for whole embryos (n p 12 for aspartic acid and glutamic acid; n p 4 for other
amino acids).
b
Values are nmol embryo21 (SEM in parentheses) for yolk-free embryos (n p 3).
804 J. E. Podrabsky and S. C. Hand
se, the proportional changes seen for most free amino acids
across development are similar whether measured in whole
embryos or yolk-free embryos (Table 4).
Because 95% of the incorporation of radioactivity into the
total free amino acid pool of these embryos can be explained
by the CPM measured in glutamic acid and aspartic acid (data
not shown), these two amino acids were used for evaluating
specific radioactivity across development. Labeling of aspartate
plus glutamate with 14C is highest in 8-dpf embryos (Tukey’s
HSD, P ! 0.05) and declines as the embryos continue to develop
and enter diapause II (Fig. 2A). The averaged specific radioactivity for these two amino acids (Fig. 2B) declines as the
embryos develop (ANOVA, P ! 0.0001 ). This decrease is a combined result of the decreased incorporation of radioactivity into
amino acids (Fig. 2A) and the increase in chemical concentration of glutamate (Table 4). The former is anticipated because
of the progressive arrest of metabolism across this period.
Rates of Protein Synthesis
Figure 2. A, Total radioactivity incorporated into aspartic acid and
glutamic acid as a function of development in embryos of Austrofundulus limnaeus. B, Specific radioactivity of aspartic acid and glutamic
acid. Embryos enter diapause II at 24 d postfertilization. Filled symbols
(mean 5 SEM, n p 4) represent developing stages, while open circles
denote diapause II stage embryos.
absence of cycloheximide (Table 2). The presence of inhibitor
causes a significant inhibition of amino acid incorporation in
all developmental stages (Table 2), and the inhibition varies
from 77.9% to 95.3% depending on the stage (Table 3). Net
incorporation of radiolabeled amino acids into acid-precipitable protein (i.e., the value without cycloheximide minus the
value with cycloheximide) increases in a linear fashion (regression analysis, P ! 0.0001 ) for all developmental stages (Fig.
1). The highest levels of incorporation are observed in the 8dpf embryos, while the lowest are found in diapause II embryos
at 42 dpf (Fig. 1).
Rates of protein synthesis (Fig. 3) were calculated for the interval between 2–4 h of incubation and were normalized to
total DNA content of the embryos to account for cellular proliferation during development (Podrabsky and Hand 1999).
The rate of protein synthesis increases from 2,033 5 374
(mean 5 SEM) at 2 dpf to 3,117 5 295 CPM mg21 DNA h21
at 8 dpf (Tukey’s HSD, P ! 0.05). Then there is a decline in
rate as the embryos continue to develop and then enter diapause II at 24 dpf (Fig. 3). Specifically, when compared with
embryos at 8 dpf, the rate of protein synthesis is depressed by
93.5% during diapause II (24 dpf; Tukey’s HSD, P ! 0.05).
Protein synthesis remains low during diapause II (340 5 53
and 203 5 53 CPM mg21 DNA h21 for 24- and 42-dpf embryos,
respectively).
In Vivo Labeling of the Amino Acid Pool
There is a 10-fold increase in total free amino acids in whole
embryos during development through diapause II (Table 4).
This total amino acid pool is dominated by the concentration
of glutamic acid, which increases dramatically during development (ANOVA, P ! 0.0001). Aspartic acid levels actually decline slightly (ANOVA, P ! 0.0001). The developmental patterns
for selected amino acids were also measured in embryos that
were dissected free of yolk before analysis (Table 4). While !1%
of the whole-embryo glutamate and only 3%–10% of the
whole-embryo aspartate are found in the embryonic cells per
Figure 3. The rate of protein synthesis in developing and diapausing
embryos of Austrofundulus limnaeus. Filled symbols (mean 5 SEM,
n p 4) represent developing stages, while open circles denote diapause
II stage embryos. Embryos enter diapause II at 24 d postfertilization.
Depression of Protein Synthesis during Diapause 805
Table 5: Cycloheximide-inhibitable oxygen consumption and ATP turnover
Oxygen Consumptiona
Days Postfertilization
8 ........................
16 .......................
24 .......................
Control
A
.182
(.006)
.111A
(.010)
.047A
(.001)
Inhibited
B
.117
(.006)
.091B
(.009)
.052A
(.003)
ATP Turnoverb
Percentage
Inhibition
Control
Inhibited
D
35.8
(1.7)
18.3
(.98)
28.8
(5.0)
.73
(.022)
.45
(.038)
.19
(.005)
.47
(.023)
.36
(.035)
.21
(.013)
.26
(.010)
.081
(.004)
2.02
(.009)
pmol O2 s21 embryo21; values are means (SEM in parentheses; n p 3). Means with the same letter are not
statistically different. Comparisons are for control versus inhibited rates for each developmental stage (paired ttest, P ! 0.002).
b
pmol ATP s21 embryo21 (assuming a P : O ratio of 2.0); values are means (SEM in parentheses; n p 3).
a
If these data are adjusted for the specific radioactivity of the
amino acid pool as measured for the intact embryo, the rate
of protein synthesis in diapause II embryos is nearly equivalent
to that in embryos at 8 dpf (data not shown). This rate of
protein synthesis cannot be supported by the measured metabolic rate of diapause II embryos (cf. the DATP turnover
caused by cycloheximide-inhibition of protein synthesis in embryos at 8 dpf with the total ATP turnover in diapause II
embryos at 24 dpf; Table 5). A further indication that this
adjustment is artifactual comes from the insignificant inhibition
of metabolism by cycloheximide during diapause, which indicates no contribution of protein synthesis to metabolism at
this stage (see below; Tables 5, 6). For these reasons, the protein
synthesis data were not adjusted for the specific radioactivity
of the free amino acid pool. Results are consistent with an exit
of cells from the cell cycle that follows an anterior to posterior
gradient along the embryonic axis as the embryos enter diapause (cf. Podrabsky and Hand 1999). Metabolically active cells
may incorporate 14CO2 into amino acids at a substantial rate,
while quiescent cells (starting in the anterior of the embryo)
may have a severely reduced rate of incorporation reflective of
a depressed metabolic state. Under this scenario, it is probable
that the specific radioactivities of amino acids in metabolically
active cells that exhibit protein synthesis are comparable across
development.
Metabolic Cost of Protein Synthesis
Cycloheximide inhibits 35.8% 5 1.7% (mean 5 SEM) of the
oxygen consumption and 36.3% 5 2.7% of the heat dissipation
in 8-dpf embryos (Tables 5, 6). The equivalent inhibition of
oxygen consumption and heat dissipation by cycloheximide
suggests that the metabolism of embryos at 8 dpf is essentially
aerobic. Assuming a P : O ratio of 2, ATP turnover attributable
to protein synthesis is calculated to be 0.26 5 0.01 pmol ATP
s21 embryo21. In contrast, statistically significant inhibition by
cycloheximide is not measurable for either oxygen consump-
tion or heat dissipation in embryos during diapause II (24 dpf),
and thus, a negligible contribution of protein synthesis to ATP
turnover in this state is indicated (Tables 5, 6).
Discussion
This study is the first to evaluate the contribution of protein
synthesis to total metabolic rate in diapausing embryos of an
annual killifish. Rates of protein synthesis are depressed by
193% in diapause II embryos of Austrofundulus limnaeus. This
decrease in protein synthesis represents a mechanism for energy
conservation that can account for 36% of the metabolic depression associated with diapause. However, arrest of protein
synthesis would presumably limit the ability of an embryo to
respond to changes in the environment via gene expression.
While not precluding limited synthesis of selected peptides,
mechanisms requiring major changes in gene expression to
facilitate survival during diapause may need to be initiated
before entry. Finally, one can infer that the half-lives of existing
proteins would be extended to facilitate long-term survival in
diapause and permit recovery from metabolic depression. Both
proteins and mRNA are known to be stabilized during anoxiainduced quiescence in embryos of Artemia franciscana (Anchordoguy et al. 1993; Hardewig et al. 1996).
Figure 4 presents rates of protein synthesis expressed per
embryo as a function of total DNA content of the embryo.
There is a strong negative relationship (r p 20.943 ) between
the amount of DNA contained in the embryo and the rate of
protein synthesis. This pattern is opposite to the expected relationship that, as DNA content increases (i.e., cell number
increases), protein synthesis should increase proportionally. Assuming that the cost of cell division and morphogenesis are
equal for the formation of structures in the anterior and posterior regions of the embryo, these data suggest that a portion
of the embryonic cells must be quiescent while others are still
active and dividing. Cycloheximide-inhibitable oxygen consumption and heat dissipation indicate that protein synthesis
806 J. E. Podrabsky and S. C. Hand
Table 6: Cycloheximide-inhibitable heat dissipation
Days Postfertilization
Controla
Cycloheximidea
Percentage
Inhibition
8 ........................
16 .......................
24 .......................
2.130 5 .006A
2.076 5 .001A
2.044 5 .005A
2.083 5 .006B
2.058 5 .001B
2.046 5 .004A
36.3 5 2.7
23.6 5 2.2
24.8 5 3.5
a
mW embryo21; values are means 5 SEM (n p 3). Means with the same letter are not
statistically different. Comparisons are for control versus inhibited rates for each developmental stage (paired t-test, P ! 0.01).
does not contribute to the metabolism of diapausing embryos
(Tables 4, 5). These data are fully concordant with our rates
of protein synthesis measured by amino acid incorporation if,
as per the above arguments, the specific radioactivity of the
amino acid pool at the site of protein synthesis is comparable
between developing and diapausing embryos.
A number of studies have shown a decrease in the rate of
protein synthesis during various states of dormancy, including
anoxia-induced quiescence (reviewed in Hand and Hardewig
1996), amphibian estivation (Fuery et al. 1998b), mammalian
hibernation (Frerichs et al. 1998), and delayed implantation in
mouse embryos (Weitlauf 1985). Further, Clegg et al. (1996)
show a substantial decrease in protein synthesis of diapausing
embryos compared with active embryos of the brine shrimp
A. franciscana. Joplin et al. (1990) report qualitative differences
in the rates of protein synthesis in brain tissue isolated from
diapausing pupae of the flesh fly, Sarcophaga crassipalpis, as
judged from a reduction in the number and intensity of protein
spots on a two-dimensional SDS-PAGE gel. Interestingly, this
group also found many continuously synthesized diapausespecific proteins, which suggests that a limited amount of protein synthesis is important in the maintenance of diapause in
Figure 4. Negative correlation (r p 20.943 ) between rates of protein
synthesis expressed per embryo and total DNA content for 8-, 16-,
24-, and 42-d postfertilization embryos. Filled symbols (mean 5
SEM, n p 4) represent developing stages, while open circles denote
diapause II stage embryos. Data for DNA content of embryos taken
from Podrabsky and Hand (1999).
S. crassipalpis (Joplin et al. 1990; Flannagan et al. 1998; Tammariello and Denlinger 1998; Yocam et al. 1998). Data presented in this study, along with those reported for diapausing
embryos of A. franciscana, delayed implanting blastocysts, and
diapausing pupae of the flesh fly indicate that depression of
protein synthesis is a common mechanism for reduction of
ATP turnover during diapause.
Depression of protein synthesis during diapause is presumably induced by endogenous cues and appears to be independent of the energy status of the cells. This is apparently the
situation in diapausing embryos of A. franciscana, which maintain high levels of adenylates and have reduced rates of protein
synthesis (Drinkwater and Crowe 1987; Drinkwater and Clegg
1991; Clegg et al. 1996). High ATP/ADP (adenosine diphosphate) ratios and adenylate energy charge also exist during
diapause II in embryos of A. limnaeus (Podrabsky and Hand
1999). However, adenosine monophosphate (AMP) levels are
elevated in diapausing embryos of A. limnaeus, and a strong
correlation has been shown between increases in the AMP/ATP
ratios and a decrease in both oxygen consumption and heat
dissipation (Podrabsky and Hand 1999). On the basis of evidence presented here, there is also a strong negative correlation
(r p 20.958) between the rate of protein synthesis and AMP/
ATP ratios during early development and diapause in A. limnaeus (Fig. 5). Lefebvre et al. (1993) found a similar correlation
between increases in AMP and decreases in the rate of protein
synthesis in hepatocytes exposed to hypoxia. Further support
for a role of AMP in the regulation of translation is found in
lysates prepared from rabbit reticulocytes (Mosca et al. 1983),
where elevated levels of AMP inhibit protein synthesis despite
the presence of high levels of ATP and GTP. These studies and
the evidence presented above suggest that AMP/ATP ratios may
be important in the regulation of protein synthesis and other
anabolic processes in diapausing embryos of A. limnaeus. The
effect of AMP could be mediated by the AMP-activated protein
kinase (AMPK; for a review see Hardie and Carling 1997; Hardie et al. 1998), perhaps through phosphorylation of key biosynthetic enzymes and initiation and elongation factors of
translation (cf. Podrabsky and Hand 1999). Mechanistic data
that link AMPK to the control of protein synthesis during
Depression of Protein Synthesis during Diapause 807
Figure 5. Negative correlation (r p 20.958 ) between the rate of protein
synthesis and AMP/ATP ratio in embryos of Austrofundulus limnaeus.
AMP/ATP ratios taken from Podrabsky and Hand (1999). Filled symbols (mean 5 SEM, n p 4) represent developing stages, while open
circles denote diapause II stage embryos.
metabolic depression are currently not available, but this possibility deserves future attention.
The arrest of protein synthesis observed in diapause embryos
of A. limnaeus can account for 36% of the metabolic depression
observed during diapause II. This value represents a substantial
energy savings to the embryo, which undoubtedly increases the
duration of developmental arrest that can be attained. Data
presented in this article and previous contributions from Clegg
et al. (1996) and Joplin et al. (1990) suggest that depression of
protein synthesis is a common mechanism for reduction of
ATP turnover during diapause. This conclusion agrees with the
accumulating body of literature concerning the role of protein
synthesis in metabolic depression during quiescence (Hand and
Hardewig 1996). A number of similarities between metabolic
depression associated with diapause and quiescence are beginning to emerge. These similarities may help elucidate mechanisms that govern large-scale changes in cellular metabolism
associated with cellular proliferation and morphogenesis.
Acknowledgments
This research was supported by a grant from the Graduate
School at the University of Colorado to J.E.P. and by NSF grant
IBN-9723746 to S.C.H.
Literature Cited
Anchordoguy T.J., G.E. Hofmann, and S.C. Hand. 1993. Extension of enzyme half-life during quiescence in Artemia
embryos. Am J Physiol 264:R85–R89.
Bradford D.F. and R.S. Seymour. 1985. Energy conservation
during the delayed-hatching period in the frog Pseudophryne
bibroni. Physiol Zool 58:491–496.
Buttgereit F. and M.D. Brand. 1995. A hierarchy of ATPconsuming processes in mammalian cells. Biochem J 312:
163–167.
Buttgereit F., M.D. Brand, and M. Muller. 1992. ConA induced
changes in energy metabolism of rat thymocytes. Biosci Rep
12:381–385.
Clegg J.S. 1976. Interrelationships between water and cellular
metabolism in Artemia cysts. V. 14CO2 incorporation. J Cell
Physiol 89:369–380.
Clegg J.S., L.E. Drinkwater, and P. Sorgeloos. 1996. The metabolic status of diapause embryos of Artemia franciscana
(SFB). Physiol Zool 69:49–66.
Darken R.S., K.L.M. Martin, and M.C. Fisher. 1998. Metabolism
during delayed hatching in terrestrial eggs of a marine fish,
the grunion Leuresthes tenuis. Physiol Zool 71:400–406.
Drinkwater L.E. and J.S. Clegg. 1991. Experimental biology of
cyst diapause. Pp. 93–117 in R.A. Browne, P. Sorgeloos, and
C.N.A. Trotman, eds. Artemia Biology. CRC, Boca Raton,
Fla.
Drinkwater L.E. and J.H. Crowe. 1987. Regulation of embryonic
diapause in Artemia: environmental and physiological signals. J Exp Zool 241:297–307.
Flannagan R.D., S.P. Tammariello, K.H. Joplin, R.A. CikraIreland, G.D. Yocum, and D.L. Denlinger. 1998. Diapausespecific gene expression in pupae of the flesh fly Sarcophaga
crassipalpis. Proc Natl Acad Sci USA 95:5616–5620.
Frerichs K.U., C.B. Smith, M. Brenner, D.J. DeGracia, G.S.
Krause, L. Marrone, T.E. Dever, and J.M. Hallenbeck. 1998.
Suppression of protein synthesis in brain during hibernation
involves inhibition of protein initiation and elongation. Proc
Natl Acad Sci USA 95:14511–14516.
Fuery C.J., P.C. Withers, and M. Guppy. 1998a. Protein synthesis in the liver of Bufo marinus: cost and contribution to
oxygen consumption. Comp Biochem Physiol 119A:
459–467.
Fuery C.J., P.C. Withers, A.A. Hobbs, and M. Guppy. 1998b.
The role of protein synthesis during metabolic depression in
the Australian desert frog Neobatrachus centralis. Comp
Biochem Physiol 119A:469–476.
Guppy M., C.J. Fuery, and J.E. Flanigan. 1994. Biochemical
principles of metabolic depression. Comp Biochem Physiol
109B:175–189.
Hand S.C. 1991. Metabolic dormancy in aquatic invertebrates.
Pp. 1–50 in R. Gilles, ed. Advances in Comparative and
Environmental Physiology. Springer, Berlin.
———. 1999. Calorimetric approaches to animal physiology
and bioenergetics. Pp. 469–510 in R.B. Kemp, ed. Handbook
of Thermal Analysis and Calorimetry. Vol. 4. Life Sciences.
Elsevier Science, Amsterdam.
Hand S.C. and I. Hardewig. 1996. Downregulation of cellular
metabolism during environmental stress: mechanisms and
implications. Annu Rev Physiol 58:539–563.
Hand S.C. and J.E. Podrabsky. 2000. Bioenergetics of diapause
808 J. E. Podrabsky and S. C. Hand
and quiescence in aquatic animals. Thermochim Acta 349:
31–42.
Hardewig I., T.J. Anchordoguy, D.L. Crawford, and S.C. Hand.
1996. Profiles of nuclear and mitochondrial encoded mRNAs
in developing and quiescent embryos of Artemia franciscana.
Mol Cell Biochem 158:139–147.
Hardie D.G. and D. Carling. 1997. The AMP-activated protein
kinase: fuel gauge of the mammalian cell? Eur J Biochem
246:259–271.
Hardie D.G., D. Carling, and M. Carlson. 1998. The AMPactivated/SNF1 protein kinase subfamily: metabolic sensors
of the eukaryotic cell? Annu Rev Biochem 67:821–855.
Hochachka P.W. and M. Guppy. 1987. Metabolic Arrest and
the Control of Biological Time. Harvard University Press,
Cambridge, Mass.
Hofmann G.E. and S.C. Hand. 1990. Arrest of cytochrome-c
oxidase synthesis coordinated with catabolic arrest in dormant Artemia embryos. Am J Physiol 258:R1184–R1191.
Joplin K.H., G.D. Yocum, and D.L. Denlinger. 1990. Diapause
specific proteins expressed by the brain during pupal diapause of the flesh fly, Sarcophaga crassipalpis. J Insect Physiol
36:775–783.
Kwast K.E. and S.C. Hand. 1993. Regulatory features of protein
synthesis in isolated mitochondria from Artemia embryos.
Am J Physiol 265:R1238–R1246.
Land S.C., L.T. Buck, and P.W. Hochachka. 1993. Response of
protein synthesis to anoxia and recovery in anoxia-tolerant
hepatocytes. Am J Physiol 265:R41–R48.
Lefebvre V.H.L., M. Van Steenbrugge, V. Beckers, M. Roberfroid, and P. Buc-Calderon. 1993. Adenine nucleotides and
inhibition of protein synthesis in isolated hepatocytes incubated under different Po2 levels. Arch Biochem Biophys
304:322–331.
Lowenstein J.M. 1967. The tricarboxylic acid cycle. Pp. 146–270
in D.M. Greenberg, ed. Metabolic Pathways. Vol. 1. Academic
Press, New York.
Mosca J.D., J.M. Wu, and R.J. Suhadolnik. 1983. Restoration
of protein synthesis in lysed rabbit reticulocytes by the enzymatic removal of adenosine 50-monophosphate with either
AMP deaminase or AMP nucleosidase. Biochemistry 22:
346–354.
Pannevis M.C. and D.F. Houlihan. 1992. The energetic cost of
protein synthesis in isolated hepatocytes of rainbow trout
(Oncorhynchus mykiss). J Comp Physiol B 162:393–400.
Podrabsky J.E. 1999. Husbandry of the annual killifish Austrofundulus limnaeus with special emphasis on collection and
rearing of embryos. Environ Biol Fishes 54:421–431.
Podrabsky J.E. and S.C. Hand. 1999. The bioenergetics of embryonic diapause in an annual killifish, Austrofundulus limnaeus. J Exp Biol 202:2567–2580.
Rolfe D.F.S. and G.C. Brown. 1997. Cellular energy utilization
and molecular origin of standard metabolic rate in mammals.
Physiol Rev 77:731–758.
SAS Institute. 1997. SAS. Version 6.0. SAS Institute, Cary, N.C.
SAS Institute. 1998. StatView. Version 5.0. SAS Institute, Cary,
N.C.
Smith R.W. and D.F. Houlihan. 1995. Protein synthesis and
oxygen consumption in fish cells. J Comp Physiol B 165:
93–101.
Storey K.B. and J.M. Storey. 1990. Metabolic rate depression
and biochemical adaptation in anaerobiosis, hibernation and
estivation. Q Rev Biol 65:145–174.
Tammariello S.P. and D.L. Denlinger. 1998. G0/G1 cell cycle
arrest in the brain of Sarcophaga crassipalpis during pupal
diapause and the expression pattern of the cell cycle regulator, proliferating cell nuclear antigen. Insect Biochem Mol
Biol 28:83–89.
Weitlauf H.M. 1985. Changes in the rate of translation with
reactivation of delayed implanting mouse embryos. J Exp
Zool 236:309–312.
Wourms J.P. 1972a. The developmental biology of annual fishes.
I. Stages in the normal development of Austrofundulus myersi
Dahl. J Exp Zool 182:143–168.
———. 1972b. The developmental biology of annual fishes.
III. Pre-embryonic and embryonic diapause of variable duration in the eggs of annual fishes. J Exp Zool 182:389–414.
Yocum G.D., K.H. Joplin, and D.L. Denlinger. 1998. Upregulation of a 23 kDa small heat shock protein transcript during
pupal diapause in the flesh fly, Sarcophaga crassipalpis. Insect
Biochem Mol Biol 28:677–682.