Myogenic cell cycle duration in Harpagifer species with sub-Antarctic and Antarctic distributions: evidence for cold compensation
1 Gatty Marine Laboratory, School of Biology, Division of Environmental and
Evolutionary Biology, University of St Andrews, St Andrews, Scotland, KY16
8LB, UK
2 Centro Austral de Investigaciones Cientificas (CADIC), Consejo Nacional de
Investigaciones Cientificas y Tecnicas (CONICET), CC92, Ushuaia, 9410, Tierra
del Fuego, Argentina
3 British Antarctic Survey, High Cross, Madingley Road, Cambridge, CB3 OET,
UK
* Author for correspondence at present address: Toxicology and Environmental Research and Consulting Laboratory, The Dow Chemical Company, 1803N Building, Midland, MI 48674, USA (e-mail: jcbrodeur{at}dow.com)
Accepted 22 December 2002
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Summary |
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Key words: cell cycle, S-phase, Harpagifer bispinis, Harpagifer antarcticus, myogenic progenitor cell, cold compensation, Antarctic, temperature, notothenioid fish, satellite cell
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Introduction |
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Modern Antarctic notothenioids are low temperature specialists possessing a
number of unique physiological adaptations to cold water that allow them to
survive at temperatures below 0°C. Some of these adaptations include the
presence of antifreeze proteins and associated aglomerular kidneys
(DeVries, 1984), a greater
conformational flexibility of proteins and enzymes
(Fields and Somero, 1998
;
Ciardiello et al., 2000
) and a
higher proportion of unsaturated phospholipid classes in cellular membranes
(Cossins, 1994
). Nevertheless,
not all physiological traits show evolutionary adjustment in Antarctic fishes.
Unloaded contraction velocity of the fast muscle fibres does not show cold
compensation in a wide range of species
(Johnston and Altringham,
1985
; Johnson and Johnston,
1991
) and maximum power output of the fast muscle is only 10-16%
of that measured in Mediterranean and tropical Perciformes
(Wakeling and Johnston, 1998
).
Similarly, comparative studies with isolated muscle mitochondria have found no
evidence for upregulation in the maximum rate of respiration per mg
mitochondrial protein in Antarctic species
(Johnston et al., 1998
), and
recent studies indicate that resting metabolic rate is not cold compensated in
polar fish (Steffensen, 2002
).
Finally, annual growth rates of the Antarctic notothenioids are also generally
believed to be slower than those of fish from temperate regions
(DeVries and Eastman, 1981
;
Everson, 1984
), although
recent analyses of growth performance data have questioned this view
(Kock, 1992
;
Kock and Everson, 1998
). These
new analyses suggest that only fish from the high-Antarctic zone, where water
temperature is stable at -1.86°C, exhibit very slow growth rates, the
growth performance of fish from lower-Antarctic latitudes/sub-Antarctic waters
being similar to that of many temperate fish species
(Kock and Everson, 1998
;
Morales-Nin et al., 2000
;
La Mesa and Vacchi, 2001
).
However, phylogenetic variation and the relatively few validations of age
estimates complicate the interpretation of this type of analysis.
Mean habitat temperature has an important influence on growth rate in
ectotherms, and the apparent slow growth of Antarctic fishes has often been
intuitively linked to the low temperatures of the Southern Ocean. However,
evidence for seasonality in Antarctic fish growth
(North, 1988;
Ashford and White, 1993
,
Coggan, 1997a
), despite the
quasi constancy of Antarctic water temperature, has led to the suggestion that
resource (i.e. food) limitation rather than temperature may restrict growth
rate in the field (Clarke,
1988
; Clarke and North,
1991
). This hypothesis is supported by laboratory experiments
which show that fish maintained under conditions characteristic of the austral
winter tend to have less appetite and consume less food than fish maintained
under summer conditions (Targett,
1987
; Coggan,
1997b
).
In fish, growth is closely linked to the dynamics of muscle growth because
the fast muscle constitutes a large proportion of the body mass
(Mommsen, 2001). The
notothenioids have an unusual pattern of muscle growth in that muscle fibre
recruitment generally ceases early in the life cycle and subsequent growth in
muscle mass is entirely by fibre hypertrophy
(Battram and Johnston, 1991
;
Fernandez et al., 2000
;
Johnston et al., 2002
). The
proliferation of a population of myogenic progenitor cells provides a source
of additional nuclei to support this process
(Koumans and Akster, 1995
;
Fauconneau and Paboeuf, 2001
).
These cells are equivalent to the satellite cells described in mammals
(Mauro, 1961
), and both
quiescent and activated progenitor cells can be identified by their expression
of c-met, the receptor for hepatocyte growth factor
(Cornelison and Wold, 1997
;
Johnston et al., 2000
), which
is believed to be involved in their activation
(Tatsumi et al., 1998
). The
proliferation of myogenic precursors is furthermore required in adult stages
for nuclear turnover and for muscle repair following injury
(Rowlerson et al., 1997
).
A crucial element of the proliferative potential of a cell population is the time needed to proceed through a complete cell cycle. Therefore, a previously unexamined way by which low temperature could restrict Antarctic fish growth rate and nuclear turnover is by limiting cell cycle progression rate in myogenic cells. The present study reports the first measurements of myogenic cell cycle duration in fish. Cell cycle duration was determined in two closely related species of Notothenioidei from the genus Harpagifer (family Notothenidae). H. antarcticus is a stenothermal species from the Antarctic peninsula that lives at temperatures between -2°C and +1°C throughout the year, while H. bispinis is a eurythermal species from the Beagle Channel, Tierra del Fuego that experiences temperatures of +4°C in winter and up to 11°C in summer. These two species have similar morphology, ecology and life history strategies, although the Antarctic species reaches a somewhat greater body size. The absence of an upward adjustment of cell cycle time at low temperatures in the Antarctic species would represent a fundamental constraint of low temperature on growth and the rate of turnover of muscle nuclei.
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Materials and methods |
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Cumulative S-phase labelling
Total cell cycle (tc) and S-phase
(ts) duration of the satellite cells were estimated using
cumulative S-phase labelling with 5-bromo-2'-deoxyuridine (BrdU; Sigma
Chemicals, Poole, UK). BrdU is a thymidine analogue that is incorporated into
DNA during the S-phase of the cell cycle. Cells labelled with BrdU can be
visualized by immunohistochemistry. The theoretical and mathematical
considerations of the cumulative S-phase labelling approach have been
described in detail by Nowakowski et al.
(1989). Briefly, the technique
involves sequentially labelling a proliferating cell population with pulses of
BrdU. As BrdU is only incorporated during the S-phase, the proportion of cells
labelled in the total population (labelling index, Li) after the first pulse
of BrdU is equal to ts/tc. With each
new pulse of BrdU, the fraction of labelled cells increases linearly as new
cells enter S-phase while previously marked cells are still visible (see
Fig. 1). This increase in Li
continues until the cells that were at the end of the S-phase at the time of
the first injection re-enter S-phase. The time needed to reach this point is
equivalent to tcts and
corresponds to the moment when Li reaches a plateau as all the proliferating
cells have been labelled (Fig.
1). The value at which the Li reaches a plateau represents the
proportion of the total population that is proliferating and is called the
`growth fraction' (GF). If the growth fraction is different from 1, the
proportion of cells marked after the first pulse of BrdU (Li0) is
more correctly expressed by:
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![]() | (2) |
![]() | (3) |
|
Experimental design
All experiments were carried out under Home Office License following
national requirements for animal care. Repeated S-phase labelling of the
satellite cells was achieved for H. antarcticus at 0°C and H.
bispinis at 5°C and 10°C by repeatedly injecting the animals
intraperitoneally with BrdU. BrdU was diluted to 10 mg ml-1 in
phosphate-buffered saline (PBS), and a volume of this solution was injected so
that the animal received 250 µg g-1 of body mass. H.
antarcticus were injected every 12 h until 132 h after the first
injection. H. bispinis were initially injected every 8 h until 72 h
and 48 h after the first injection for fish at 5°C and 10°C,
respectively. However, as more time was needed for the labelling index to
reach a plateau, a second group of H. bispinis was injected every 12
h until 144 h and 120 h after the first injection for fish at 5°C and
10°C, respectively. To monitor the evolution of the labelling index and
establish a curve like the one shown in
Fig. 1, one animal from each
group was killed by a sharp blow to the head 1 h after each injection. Fish
were, however, not sampled after the first injection because the number of
labelled cells was usually too low to be accurately measured at that time. A
5-mm thick transverse slice of the trunk was taken at 0.7 fork length. The
slices were frozen on cork strips in isopentane cooled near its freezing point
in a mixture of acetone and dry ice and were stored at -20°C until
sectioning. Frozen trunk slices were cut on a cryostat at 7-µm thickness
and mounted on glass slides coated with poly-L-lysine. The sections were
air-dried and stored at -20°C until processing for
immunohistochemistry.
Immunohistochemistry
Muscle sections were doubly stained against BrdU and c-met to localize
labelled myogenic cells (Fig.
2). The primary antibodies used were a rabbit polyclonal antibody
against c-met (m-met, SP260; Santa Cruz Biotechnology, Santa Cruz, USA) and a
mouse monoclonal antibody against BrdU (Clone BU-33, Sigma Chemicals).
Sections were first fixed in acetone for 10 min and then placed in a solution
containing 5% (v/v) normal goat's serum, 1% (v/v) Triton X-100 and 1% (m/v)
bovine serum albumin (BSA) in PBS for 15 min to rehydrate them and block
non-specific binding sites. Sections were subsequently washed for 3x2
min in PBS and incubated overnight at 4°C with both primary antibodies
diluted 1:20 in a solution containing 1% Triton X-100 and 1% BSA in PBS. After
three washes in PBS, sections were incubated for 30 min at room temperature
with biotinylated goat anti-mouse secondary antibody (Sigma Chemicals) diluted
1:20 in a preparation of secondary anti-rabbit antibody linked by a dextran
polymer to a number of horseradish peroxidase molecules obtained from DAKO A/S
(DAKO EnVisionTM; Ely, UK). The sections were again washed for 3x2
min in PBS and incubated for 30 min at room temperature with alkaline
phosphatase-conjugated extraAvidin (Sigma Chemicals, Poole, UK) diluted 1:20
in a solution containing 1% Triton X-100 and 1% BSA in PBS. After washing the
sections for 3x2 min in Tris buffer, alkaline phosphatase activity was
developed using a solution containing Fast Blue BB, levamisole,
naphtol-ASMX-phosphate and N,N-dimethylformamide in Tris buffer,
which gives a blue end product (Van der
Loos, 1999). Peroxidase activity was afterward developed using
3-amino-9-ethylcarbazole, which gives a red end product
(Van der Loos, 1999
). Counts
of the number of red (c-met positive), blue (BrdU positive) or purple (c-met
and BrdU positive) cells were made from 60 fields of 0.024-0.031
mm2 per fish.
|
Statistical analysis
The increase of the labelling index over time was modelled at each
temperature by linear regression analysis. The slopes of the regression lines
fitted at each temperature were compared by analysis of covariance using the
general lineal model (GLM) of the SPSS statistical software.
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Results |
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Total cell cycle and S-phase duration were determined by solving equations 2 and 3 (see Materials and methods). Both cell cycle and S-phase duration were longer at lower temperatures in H. bispinis (Table 2), with a Q10 for cell cycle progression of 3.4. Using this Q10, a cell cycle time of 277 h was predicted for H. bispinis at 0°C, which is much more than the cell cycle time of 111 h measured in H. antarcticus at this temperature (Table 2).
|
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Discussion |
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The present study showed that cell cycle duration increases with decreasing temperatures in H. bispinis (Table 2). However, in contrast with this finding, cell cycle time was 1.35 times shorter in the Antarctic H. antarcticus at 0°C than in the sub-Antarctic species at 5°C (Table 2), suggesting a cold compensation of cell cycle progression rate in the Antarctic species. Two features of the data obtained suggest that this adjustment of satellite cell cycle duration in H. antarcticus represents an evolutionary adaptation. Firstly, the adjustment is in the direction to elevate cell cycle time in the polar species when compared with the sub-Antarctic species. Secondly, the cell cycle time of the two species is almost identical when compared at the median annual habitat temperature at which each species lives (H. antarcticus 111 h at 0°C and H. bispinis 110 h at its median life temperature of 7.5°C, estimated by interpolation using the observed Q10 of 3.4). Taken together, the data obtained in this study therefore suggest that evolution has been able to adjust the basic molecular machinery of cell proliferation to offset the rate-limiting effect of low temperatures.
Comparative data on myogenic cell cycle time in vivo are confined
to a single study on 30-day-old rats
(Schultz, 1996). This study
found that 80% of fast skeletal muscle satellite cells proliferated with a
cell cycle duration of 32 h and an S-phase of 14 h
(Schultz, 1996
). This
percentage of proliferating satellite cells is similar to the 75% measured in
the two Hapagiferidae species. However, whereas Schultz
(1996
) suggested that the
remaining 20% of satellite cells were also dividing, although at a slower
rate, the presence of linear labelling and a distinct plateau in
Fig. 3 suggests a more
homogeneous cell cycle time in the present study
(Nowakowski et al., 1989
). The
relatively high proportions of dividing myogenic cells measured in both rats
and fish may be associated with nuclear turnover as well as growth, since
studies in the rat have shown that 2% of the nuclei are replaced each week in
adult stages (Schmalbruch and Lewis,
2000
). The cell cycle times measured in the present study probably
reflected this nuclear turnover since the Harpagifer used were not
actively growing.
Numerous extracellular signals that influence growth rate (e.g. growth
factors, mitogen antagonists, differentiating inducing factors) are involved
in regulating the cell cycle (Beijersbergen
and Bernards, 1996; Jones and
Kazlauskas, 2001
). In the present study, the Harpagifer
species had growth rates that were not significantly different from zero. The
observed differences in cell cycle time are therefore most likely due to
differences in temperature. Although the specific molecular and physiological
adaptations responsible for the cold compensation of cell cycle progression
rate remain to be identified, it is likely that already described cold
adaptations of Antarctic fish such as the structural alteration of tubulins
(Dietrich, 1997
) and the
greater conformational flexibility of proteins and enzymes
(Fields and Somero, 1998
;
Ciardiello et al., 2000
) play a
part in this phenomenon. Interestingly, whereas total cell cycle time was
considerably shorter in H. antarcticus at 0°C than in H.
bispinis at 5°C, the duration of the S-phase was similar between the
two species (Table 2),
indicating that cold compensation was less important for DNA replication rates
than for other parts of the cell cycle.
Although it is still debated whether or not growth is slower in Antarctic
fishes, the present study shows that, if such a restriction of growth rate
exists, it is unlikely to be due to a limitation of muscle growth by low
temperature since myogenic cell cycle progression rate appears to show cold
compensation. However, it remains to be examined whether cell cycle
progression rate also shows cold compensation in myogenic progenitors of
high-Antarctic fishes, as there are indications that these fish may be the
only notothenioids with a truly reduced growth rate
(Kock and Everson, 1998;
Morales-Nin et al., 2000
;
La Mesa and Vacchi, 2001
).
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Acknowledgments |
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