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First published online April 18, 2008
Journal of Experimental Biology 211, 1482-1489 (2008)
Published by The Company of Biologists 2008
doi: 10.1242/jeb.015636
Functional and morphological plasticity of crocodile (Crocodylus porosus) salt glands
School of Integrative Biology, The University of Queensland, St Lucia, Brisbane, 4072 Australia
* Author for correspondence (e-mail: c.franklin{at}uq.edu.au)
Accepted 10 March 2008
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Summary |
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 TOP Summary INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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Key words: osmoregulation, reptile, tissue respirometry, stereology, transmission electron microscopy
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INTRODUCTION |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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to over 60 (Grigg
et al., 1986
). Data from field surveys demonstrate that animals
can maintain plasma osmolarity within a narrow range (298–309 mOsm)
along such salinity gradients (Grigg,
1981). In order to successfully osmoregulate in saline environs,
C. porosus utilise extra-renal salt-secreting glands
(Taplin and Grigg, 1981), much
like marine reptiles and pelagic sea birds. Marine birds have paired orbital
salt glands, which secrete excess salt
(Fänge et al., 1958;
Schmidt-Nielsen et al., 1958),
whereas in marine turtles, lachrymal salt glands are responsible for salt
secretion (Schmidt-Nielsen and Fänge,
1958). Sea snakes, however, have sublingual salt glands, which
open into the oral cavity (Dunson and
Taub, 1967). In crocodiles, the salt-excreting glands are located
on the tongue immediately below the lingual epithelium and are responsible for
the excretion of sodium and chloride ions
(Taplin and Grigg, 1981;
Taplin et al., 1982).
Lingual salt glands, unique to crocodilians, were first identified and
described in C. porosus by Taplin and Grigg
(Taplin and Grigg, 1981). The
salt glands appear as 20–40 distinct pores on the posterior half of the
lingual surface. Classified as compound, branched, tubular structures, each
gland is composed of 10–14 distinct lobes. Lobes consist of a number of
short, blind-ending secretory tubules which join together to form multiple
interlobular ducts. Secretions exit the gland via these ducts.
Numerous arterioles and venules punctuate the connective tissue around and
between gland lobes, and a dense network of capillaries can be found in the
connective tissues both surrounding and between the secretory tubules
(Franklin and Grigg, 1993).
Nerve fibres are also present in large numbers in the connective tissues
between and throughout the salt gland lobes and tubules
(Cramp et al., 2007;
Franklin et al., 2005).
Methacholine chloride, an acetylcholine agonist, stimulates lingual salt gland
secretion in C. porosus (Taplin
and Grigg, 1981) suggesting that active secretion may be under the
control of the cholinergic nervous system
(Franklin et al., 2005). In
marine birds and turtles, salt loading by intravenous or intraperitoneal
infusion of hypertonic salt solutions also stimulates secretions from the salt
glands (e.g. Fänge et al.,
1958; Schmidt-Nielsen and
Fänge, 1958;
Schmidt-Nielsen et al., 1958).
Salt loading of restrained crocodiles does not stimulate secretion from the
salt glands; an increase in the rate of secretion from the salt glands
following salt loading only occurs in those that are freely moving and
unstressed, implying a significant involvement of the adrenergic nervous
system in regulating (suppressing) crocodilian salt gland secretion
(Franklin et al., 2005;
Taylor et al., 1995).
Avian salt glands show remarkable levels of phenotypic plasticity –
both morphological and physiological – as a consequence of drinking
saline water (for a review, see
Shuttleworth and Hildebrandt,
1999). In birds that have never been exposed to saline conditions,
salt glands secrete little; initial exposure to an osmotic stressor initiates
a cascade of morphological and physiological responses, which results in a
marked increase in salt gland secretion rate. The most striking change in
avian salt glands upon initial exposure to salt water is the increase in gland
size (Ellis et al., 1963)
which is brought about by changes in secretory cell volume, cell number and
blood flow to and within the gland
(Ballantyne and Wood, 1969;
Hanwell et al., 1971;
Hanwell and Peaker, 1975;
Hossler, 1982;
Knight and Peaker, 1979).
These changes in morphology accompany increases in mitochondrial and
glycolytic enzymes (McFarland et al.,
1965; Spannhof and Jurss,
1967; Stainer et al.,
1970) and increases in the abundance and activity of
Na+,K+-ATPase (e.g.
Ernst et al., 1967;
Ernst and Mills, 1977;
Fletcher et al., 1967;
Stewart et al., 1976). In
addition, most of these adaptive responses appear to be reversible during
de-adaptation (e.g. Hossler et al.,
1978; Mcarthur and Gorman,
1978).
Given the wide range of salinities across which the estuarine crocodile
Crocodylus porosus can successfully osmoregulate in the wild, we
speculate that crocodile salt glands should demonstrate a broad functional
flexibility. Moreover, this functional flexibility should also be manifested
in the cellular morphology of the glands. In previous studies, it has been
shown that the gross morphology of the salt glands changes significantly with
prolonged acclimation to saline environments
(Cramp et al., 2007;
Franklin and Grigg, 1993).
Although we know that C. porosus transferred from fresh to salt water
are immediately capable of maintaining the plasma ion homeostasis
(Taplin, 1985), suggesting
that salt gland function can be upregulated rapidly, little is known about
long-term acclimatory changes in the structure and function of the glands
following prolonged exposure to hypersaline environs.
In the present study, we have examined whole animal osmoregulatory capacities, maximal secretory rates, and tissue metabolic rate in C. porosus salt glands following prolonged acclimation to both fresh and hypersaline environments. Additionally, we performed a detailed morphological study of salt gland secretory cells from acclimated animals to determine if morphological changes to cells occur with prolonged saltwater acclimation which may correlate with functional changes in the tissue.
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MATERIALS AND METHODS |
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2 weeks), animals were assigned to salinity treatment
groups. Hatchling crocodiles were randomly assigned to one of two treatment groups: freshwater (FW) or saltwater (SW) acclimation. SW-acclimated animals were placed into 70% seawater made using commercially available artificial sea salt (Aquasonic, Wauchape, New South Wales, Australia). Animals in both treatments were fed and had their water changed twice weekly. Animals remained in these treatments for 6 months.
Maximal secretory rate
Animals (N=6 per treatment group) were fasted for 1 week prior to
experimentation. Animals were removed from holding tanks, weighed and
restrained on a foam-lined board and their mouths propped open with a cork and
secured with fabric tape. The tongue surface was rinsed clean with water and
then blotted dry. Animals then received 2 mg kg–1
methacholine chloride in sterile saline via intraperitoneal injection
and were left undisturbed for 10 min. Whatman filter papers pieces were cut to
the size and shape of the animal's tongue. Secretions from the salt gland
pores were collected by placing the filter papers over the tongue and pores
and allowing the secretion to be absorbed into the filter paper. Filter paper
pieces were large enough that they did not become completely saturated with
secretion during the sampling period and that ensured secretions were
continually absorbed by capillary action into the paper. Filter paper pieces
were left on the tongue for 10 min. At the end of 10 min, the filter paper was
removed and placed into a glass vial containing 1 ml of ultrapure water
(Millipore, 18.2 M
cm). The process of secretion collection was
repeated three more times. Vials containing salt secretions were allowed to
sit overnight for the salt to diffuse into the surrounding fluid.
Na+ concentrations were measured with an EEL flame photometer
(Corning Glassworks, Corning, NY, USA), taking into account the additional
Na+ present in the filter paper prior to the addition of the
secretions. Na+ secretion rate data were scaled to body mass and
are presented as µmol 100 g–0.7 body mass
h–1. The highest rate of secretion achieved over the 40 min
period was deemed the maximal secretory rate. Mass and maximal secretory rates
were compared with Student's t-tests using the statistical program
SigmaStatTM. Data are presented as means ± s.e.m.
Blood collection and processing
A 1 ml blood sample was taken from the cervical sinus of six crocodiles
from each treatment group into individual heparinised syringes. Form these,
duplicate 50 µl samples were taken and placed in haematocrit tubes and
centrifuged for 2 min at 13 000 g. The remaining blood sample
was centrifuged at 2500 g for 3 min and the plasma collected.
Osmotic pressure was measured by vapour pressure osmometry, and
Na+, K+ and Cl– concentrations were
determined using an Olympus AU400 automated analyser (Olympus America Inc.,
Melville, NY, USA) using the ion-selective electrode (ISE) method. All
haematological parameters were compared with Student's t-tests or
where data failed tests of normality or equal variance, non-parametric
Mann–Whitney rank sum tests were performed. All analyses were performed
with the statistical program SigmaStatTM. Data are presented as means
± s.e.m.
Tissue respirometry
In vitro salt gland oxygen consumption rate was measured using a
Strathkelvin Instruments Mitocell S200 micro respirometry system comprising a
782 oxygen meter, 1302 oxygen electrode MT200 respirometer (Strathkelvin
Instruments, Motherwell, Scotland). Crocodiles (N=5 per treatment)
were killed by overdose of anaesthetic (sodium thiopentone, Jurox, Rutherford,
New South Wales, Australia). The tongue was excised whole and placed into
ice-cold crocodile ringer (containing in mmol l–1, NaCl 116,
KCl 4, CaCl2 2, MgCl2 2, glucose 15, sodium pyruvate 5,
Hepes-Na 10). The salt glands were dissected free of the underlying connective
tissue and overlying epithelia. Small (<0.5 mm thick
1 mm2)
slices of salt gland tissue were placed into beakers containing ringer at room
temperature bubbled continuously with air and allowed to incubate for 15 min.
One piece was then placed into a MT200 microrespirometer containing a magnetic
stirrer bar and 0.5 ml of air-saturated crocodile ringer at 25°C. The
chamber was sealed and the rate of oxygen consumption by the tissue in the
chamber was recorded instantaneously until the partial pressure of oxygen in
the chamber had dropped by 20% (approximately 30 min, depending on the
treatment group). At the end of this period the chamber was opened and the
ringer was replaced with fresh oxygenated ringer containing 0.1 mmol
l–1 methacholine chloride. The chamber was once again sealed
and the rate of oxygen consumption measured until the partial pressure of
oxygen in the chamber had dropped by 20%. Prior to the commencement of
experimentation, trials were conducted to ensure that oxygen consumption rates
did not drop substantially over the course of the experiment. The data (not
shown) indicated that, apart from an immediate drop in the metabolic rate of
the tissue following excision (first 15 min), the metabolic rate of the tissue
was subsequently stable for at least 4 h. Care was taken to ensure that all
experimental measures were completed within this 4 h window. At the completion
of the experiment, tissue pieces were blotted dry and weighed. Prior to the
commencement of, and at the end of each trial, the rate of background oxygen
consumption in the chamber (from contaminants and the electrode) in the
absence of any tissue was recorded and this mean rate subtracted from the
tissue rates. In order to minimize bacterial contamination of the chamber,
chambers were cleaned prior to trials with 70% ethanol and the chamber ringer
was filtered through 0.2 µm Millipore syringe filters. Rates of oxygen
consumption were standardized to tissue wet mass and are reported as µl
O2 g–1 h–1. Data were compared
statistically using a two-way repeated-measures AVOVA. Post-hoc
comparisons were made with Holm–Sidak tests. All statistical comparisons
were made using the statistical program SigmaStatTM. Data are presented
as means ± s.e.m. and treatments were considered as significantly
different if P<0.05.
Transmission electron microscopy
Crocodiles (N=4 per treatment) were euthanased and salt glands
dissected out as described previously. Small pieces (circa 2
mm2) containing at least one visible pore were dissected out from
the surface layer and were placed into cold 2% glutaraldehyde (ProSciTech,
Thuringowa, Australia) in 0.1 mol l–1 phosphate buffer pH 7.4
for 24 h. Following glutaraldehyde fixation, tissues were washed in phosphate
buffer and then post-fixed in 1% osmium tetroxide (ProSciTech, Thuringowa,
Australia) in phosphate buffer. Tissue pieces were then dehydrated through an
ascending ethanol gradient and infiltrated with Spurrs medium (ProSciTech).
Tissues were blocked and allowed to polymerise at 60°C for 48 h. Semi-thin
sections were cut, stained with 1% Toluidine Blue in 0.1 mol
l–1 phosphate buffer and viewed under a microscope to
determine if the appropriate position in the tissue piece had been reached
(i.e. there were secretory cells present). Once the appropriate position
within the tissue block was confirmed, ultrathin sections (60 nm) were cut
with a diamond knife and collected onto 2 mm copper grids. All sectioning was
performed with a Leica Ultracut UC6 Ultramicrotome. Sections were stained with
uranyl acetate and lead citrate and viewed at 80 kV in a Jeol 1011
transmission electron microscope. Images were captured with a MegaView III
digital camera using the iTEMTM software package.
Stereological analysis
Between 50 and 70 images of different secretory cell were collected by
transmission electron microscopy (TEM) from four animals per treatment group.
Mitochondrial volume density, plasma membrane surface area, intercellular
space volume, and mitochondrial cross-sectional area were measured using the
morphometrics software package, SigmaScanProTM.
Volume density of mitochondria within secretory tissue (VV mit)
The volume density of mitochondria within glandular secretory tissue
(VV mit) was determined by systematic point counting of
mitochondria in ultrathin sections. Using image analysis software, a coherent
test lattice (30x24 µm; with lines spaced 2 µm apart) was
superimposed over digital transmission electron micrographs of glandular
tissue. The number of points of intersection (i.e. where test lines cross one
another) over mitochondria was then counted. To calculate VV
mit, this number was divided by the total number of points of
intersection over secretory tissue.
Volume density of intercellular space within secretory tissue (VV ics)
The volume density of intercellular space within glandular secretory tissue
(VV ics) was determined by systematic point counting using
the same coherent test lattice and digital TEMs of glandular tissue as used to
determine VV mit. To determine intercellular space, the
number of points of intersection over areas of intercellular space was counted
and divided by the total number overlaying secretory tissue.
Surface density of basolateral cell membrane within secretory tissue (SD cm)
The surface density of basolateral cell membrane within secretory tissue
(SD cm) was determined using a coherent test lattice
(30x24 µm; with lines spaced 4 µm apart) superimposed over the
same TEMs used to determine VV mit and VV
ics. To determine SD cm, the number of times test
lines intersected the basolateral membrane of secretory cells was counted in
micrographs from each animal. To calculate SD cm, this number was
doubled and then divided by the total length of line overlaying secretory
tissue (LC). LC was calculated using
the equation:
LC=LTxVV st,
where LT is the total length of test line applied and
VV st is the volume density of secretory tissue (st) in
micrographs.
Cross-sectional area of mitochondria (µm2)
Differences in VV mit between treatment groups could
reflect a difference in the size and/or density of mitochondria. To
investigate differences in VV mit, we compared
mitochondrion cross-sectional area between treatment groups. We did this by
measuring the cross-sectional area of 200 mitochondria from each animal in
both treatment groups. Cross-sectional area was measured directly from digital
TEMs of glandular tissue using the image analysis program (SigmaPlotTM
7.0).
Cell density
SD cm will depend not only on the extent of elaboration of the
basolateral cell membrane, but also the number of cells per unit volume of
secretory tissue. To investigate the effects of saltwater and freshwater
acclimation on basolateral cell membrane area more thoroughly, we compared the
number of nucleated cells, per unit volume density of secretory tissue
(VV st), in micrographs from each animal. We did this by
counting the number of nucleated cells in TEM micrographs and dividing this by
the volume density (VV st) of glandular secretory tissue.
All morphological parameters were compared with Student's t-tests
using the statistical program SigmaStatTM. Data are presented as means
± s.e.m.
Chemicals
All reagents were sourced from Sigma Aldrich (Sydney, Australia) unless
otherwise specified.
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RESULTS |
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Ultrastructure of principal secretory cells
The secretory tubules of C. porosus consist of a single layer of
simple cuboido-columnar epithelial cells. The epithelium is composed primarily
of just one cell type, the principal secretory cell, although this cell type
appeared to have several morphological states. At the blind end of the
secretory tubules, the principal cells were typically conical in cross
section. They were, on average approximately 15 µm long and 7 µm wide at
their widest point, tapering to approximately 3 µm at the apical tip of the
cell (Fig. 4A).
Morphologically, the principal cells were characterised by a greatly amplified
basolateral membrane (BLM). Lateral, microvilli-like projections of the BLM
intermeshed with those of adjacent cells
(Fig. 4B). By contrast, the
apical membrane surface area was relatively reduced. A few, small microvilli
projected from the apical surface of the secretory cells into the secretory
tubule lumen (Fig. 4C).
Adjacent principal cells were linked at their apexes by zona occludens and
zona adherens junctions and along their basolateral membranes by zona adherens
junctions. Principal cells contained a large, basally located nucleus composed
primarily of euchromatin, with small regions of heterochromatin at the nuclear
periphery. The cells generally contained a large number of mitochondria, which
were evenly dispersed throughout the cell. Numerous secretory vesicles were
readily visible at the apical tip and, to a lesser degree, along the
basolateral membrane of the principal cells. Large amounts of smooth
endoplasmic reticulum (ER) could be seen throughout the cell. Rough ER was
less evident in principal cells. In cross section most cells could be seen to
contain at least one Golgi complex, generally located close to the nucleus.
Many secretory cells also contained small electron-opaque, non-membrane bound
vesicles. Principal cells rested upon a well-developed basement membrane and
were in close proximity to capillaries
(Fig. 4D).
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Morphological plasticity of principal secretory cells
There was no significant effect of saltwater acclimation on either the
surface density of the basolateral cell membrane within secretory tissue
(SD cm; Table
1; t-test, P=0.714, d.f.=7) or the volume of
intercellular space between secretory cells (VV ics;
Table 1; t-test,
P=0.546, d.f.=7). There was a significant effect of saltwater
acclimation on the volume density of mitochondria within the principal
secretory cells; there was an increase of almost 23% in mitochondrial volume
per µm3 in SW-acclimated animals relative to FW-acclimated
animals (Table 1;
t-test, P=0.024, d.f.=7).
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There were no differences in mitochondrial cross-sectional areas between FW- and SW-acclimated animals (t-test, P=0.55, d.f.=7), suggesting that the increase in mitochondrial volume density is most probably due to an increase in mitochondrial numbers, rather than mitochondrial size.
There was a significant difference in the numbers of cell nuclei per µm2 per micrograph, with SW-acclimated animals having significantly fewer nuclei than FW-acclimated animals (Table 1; t-test, P=0.046, d.f.=7), suggesting that cells from SW-acclimated animals were larger than those of FW-acclimated animals.
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DISCUSSION |
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;
Wells et al., 1991),
suggesting that SW-acclimated crocodiles were able to successfully maintain
osmotic homeostasis despite extended exposure to hyperosmotic conditions.
Moreover, extended exposure to hypersaline conditions appeared to have little
long-term impact on the animal's growth, suggesting that the additional energy
required in maintaining homeostasis was either relatively minimal or was
compensated for by increased food intake (animals were fed ad
libitum, so we cannot readily distinguish between these
possibilities).
Surprisingly, blood haematocrit, plasma osmolarity and plasma
Na+ and K+ concentrations were significantly lower in
FW-acclimated crocodiles when compared with their SW-acclimated counterparts.
In previous studies of plasma Na+ concentrations and water flux in
C. porosus, Taplin (Taplin,
1985) demonstrated that FW-acclimated C. porosus actively
drank freshwater when available, but when living in hyperosmotic environments,
animals avoided drinking saltwater and obtain all necessary water though
ingested food and metabolic water (Taplin,
1985). The active consumption of freshwater by C.
porosus, when available, is thought to be related to the method of
excretion of nitrogenous waste utilised while in freshwater [i.e.
predominantly ammonia in freshwater as opposed to insoluble urates when in
saltwater (Grigg, 1981)] and
the associated high requirement for water. The ingestion of such an
electrolyte-depleted medium, whilst important for waste excretion, may
secondarily be responsible for the relatively reduced plasma osmolarity and
haematocrit, and lower electrolyte levels in FW-acclimated animals observed in
this study. Moreover, the integument of the cephalic region (including the
buccal cavity, but excluding the salt glands) is also highly permeable to
Na+ and is a major secondary route for Na+ efflux (and
possibly other ions) in C. porosus
(Taplin, 1985). Hence, in
freshwater, maintaining a lower plasma osmolarity, relative to SW-acclimated
animals, may aid in reducing the rate of Na+ efflux from the buccal
integument into the surrounding medium.
To investigate salt gland functional capacity in vivo,
methacholine chloride, an acetylcholine (ACh) agonist, was used to stimulate
maximal salt gland secretory rates in restrained C. porosus.
Previously, Taplin reported that short term exposure (160 h) to saline
environments in C. porosus hatchlings reared in freshwater was not
long enough to induce differences in maximal salt gland secretion rates
relative to animals maintained continuously in freshwater
(Taplin, 1985). However, in
this study we found that chronic exposure to hypersaline environments resulted
in adaptive changes to the salt gland, which enabled a significantly elevated
rate of Na+ secretion. Maximal mean lingual gland secretory rates
were more than three times higher in SW-acclimated C. porosus
relative to FW-acclimated animals. In FW-acclimated C. porosus,
methacholine stimulation appeared to have relatively little effect on
secretion rate. These results possibly suggest that prolonged saltwater
acclimation results in changes to the cholinergic pathways responsible for
salt secretion.
Although acetylcholine can act on numerous aspects of the tissue to
increase secretion rate, we believe that there is at least an underlying
direct increase in the response of the secretory tissue to exogenous
methacholine, which may increase the secretory capacity of the tissue. In
SW-acclimated C. porosus salt gland tissue slices, application of
methacholine had a marked effect on the in vitro metabolic rate of
the tissue, raising it by over 30%. A comparable metabolic response to
methacholine application, in vitro, has been observed in both avian
and turtle salt gland slices (Shuttleworth
and Thompson, 1987). In contrast to tissue slices from
SW-acclimated crocodiles, those from FW-acclimated animals showed relatively
little response to methacholine. This result suggests that tissue slices from
SW-acclimated crocodiles are more responsive to ACh than those of
FW-acclimated crocodiles.
It was recently demonstrated that crocodile salt glands are richly
innervated by cholinergic neurons, suggesting a large role for ACh in the
regulation of salt gland secretion
(Franklin et al., 2005).
Similarly, avian salt-secreting tissues also appear to be under the control of
the cholinergic system (e.g. Ash et al.,
1969; Schmidt-Nielsen and
Fänge, 1958). Moreover, avian salt gland secretory cells also
possess large numbers of muscarinic ACh (mACh) receptors, which when
activated, initiate a multitude of secretion-related processes within the cell
(e.g. Borut and Schmidt-Nielsen,
1963; Hootman and Ernst,
1981; Hootman and Ernst,
1982; Stewart et al.,
1979; van Rossum and Ernst,
1978). In addition, saltwater acclimation results in a 3-fold
increase in the number of mACh receptors on duck salt gland secretory cells
(Hootman and Ernst, 1981),
suggesting that ACh plays a large, multifaceted role in regulating the
secretory process. We suspect that, as in avian models, ACh is likely to act
directly on the secretory cells in the salt glands of C. porosus to
regulate the secretory activity of the tissue. Moreover, saltwater acclimation
is likely to result in an increase in ACh receptor numbers in C.
porosus secretory cells, although this, and the other potential sites of
action by ACh, remains to be empirically determined.
Increases in the maximal secretory capacity of SW-acclimated crocodiles
were reflected by changes in secretory tissue morphology. Studies on birds and
turtles have demonstrated that salt-secreting tissues are morphologically
plastic and are capable of responding rapidly to changes in plasma osmolarity
(e.g. Abel and Ellis, 1966;
Benson and Phillips, 1964;
Ernst and Ellis, 1969;
Reina, 2000;
Schmidt-Nielsen and Kim,
1964). Changes include increases in blood flow to the gland
(Hanwell et al., 1970) and the
rapid increase in salt gland size by both cellular hypertrophy and hyperplasia
(Abel and Ellis, 1966;
Benson and Phillips, 1964;
Reina, 2000;
Schmidt-Nielsen and Kim,
1964). In addition, changes in secretory cell ultrastructure,
particularly marked increases in plasma membrane surface area and increases in
the number of mitochondria (Abel and Ellis,
1966; Bokenes and Mercer,
1998) accompany increases in gland size. In C. porosus we
found no increase in the surface density of plasma membrane per unit area in
SW-acclimated animals, but did observe an increase in the volume density of
mitochondria. As mitochondrial cross-sectional areas were not different across
treatments, it suggests that changes in secretory cell mitochondrial volume
per unit area arise from an increase in mitochondrial numbers (density per
unit volume) in SW-acclimated crocodiles. Mitochondrial density is an
important proxy for estimates of the metabolic activity of a cell. Hence cells
involved in energetically expensive processes such as secretion and absorption
often have large numbers of mitochondria. Salt-secreting cells generally have
relatively large numbers of mitochondria. Indeed, actively secreting avian
salt gland cells have approximately twice the volume of mitochondria compared
with their inactive counterparts (Bokenes
and Mercer, 1998). Moreover, the metabolic rate of secretory cells
from SW-acclimated ducks is almost double that of naive (FW-acclimated)
animals (Hootman and Ernst,
1980).
In the present study, we also examined the rate of oxygen consumption by
isolated crocodile salt gland tissue and found there was no difference in the
mass-specific metabolic rate of tissues from FW- and SW-acclimated crocodiles.
There are fewer cells per unit mass in SW-acclimated salt gland relative to
FW-acclimated ones (Cramp et al.,
2007) and when oxygen consumption rates are expressed in terms of
the number of cells per unit mass, the metabolic rate of SW-acclimated
crocodile salt gland tissue would be higher than that of FW-acclimated cells.
Similar values are seen in duck salt gland models
(Hootman and Ernst, 1980).
Given that there are almost 23% more mitochondria per unit volume in
SW-acclimated crocodiles, and again, since these secretory cells are so much
larger than those of FW-acclimated crocodiles, this figure is also probably an
underestimate of the total cellular volume of mitochondria.
Following prolonged saltwater acclimation, we observed no increase in the
density of plasma membrane per unit volume of salt gland tissue relative to
that from FW-acclimated crocodiles. As detailed above, this result is likely
to be an underestimate of the actual value since there are fewer secretory
cells per unit volume of SW-acclimated salt gland
(Cramp et al., 2007). We
suspect that when the surface area measurements are expressed relative to
total cells size, there would be an increase in plasma membrane surface area
of salt gland following acclimation to hypersaline environments. However, the
extent of the change in plasma membrane area from SW-acclimated crocodile
cells is significantly less than is observed in duck salt gland cells
following osmotic challenge, in which there can be a fivefold increase in
plasma membrane surface area (Merchant et
al., 1985). These differences in plasma membrane response to salt
loading may simply reflect species differences between crocodiles and ducks,
but may reflect greater underlying differences in their abilities to respond
to osmotic challenges over long periods of time. Further work is required to
determine whether the ultrastructural changes we observed in chronically
SW-acclimated salt glands are different to those that occur following acute
osmotic challenge.
Relative to the salt glands of birds, the functional and morphological
effects of saltwater acclimation in C. porosus were comparatively
small. Although it is possible that these differences reflect different time
courses over which these responses occurred, it is also possible that
differing relative salt loads may also play a role. C. porosus can
successfully obtain all necessary water requirements from ingested food and
metabolic water production alone, without the need to maintain water balance
by drinking seawater (Taplin,
1984; Taplin,
1985). Indeed, several species of marine reptiles (e.g. sea snake
Pelarmis platurus, estuarine turtles Callagur borneoensis
and Malaclemys terrapin and Crocodylus acutus) are thought
to also maintain water balance without the need to drink seawater
(Dunson, 1982;
Dunson and Moll, 1980;
Dunson and Robinson, 1976;
Evans and Ellis, 1977;
Robinson and Dunson, 1976).
Taplin (Taplin, 1985) provided
data on the relative salt loads of saline water food items of C.
porosus in the Liverpool/Tomkinson River System, northern Australia,
which showed that the Na+ content of these items ranged from
55 to 215 mmol Na+ kg–1 body mass (the
mean Na+ content of food items fed to crocodiles in the present
study was 145 mmol kg–1 body mass). Hence when living in
seawater, a crocodile's salt load would be equivalent to that of the food it
consumes plus any incidental water ingested (i.e. during swallowing). By
contrast, marine birds maintain hydration by consuming seawater and excreting
the excess salt through their salt glands. With an average [Na+] of
around 450 mmol l–1, the ingestion of seawater would
contribute to a much greater relative salt load than that experienced by
crocodiles. Hence, differences in acclimatory responses to seawater exposure
between marine birds and elasmobranchs and crocodiles may reflect different
relative salt loads. We would hypothesise that increases in dietary salt loads
may stimulate a great phenotypic response in the salt glands of C.
porosus.
Crocodylus porosus has a remarkable ability to successfully
inhabit severely hypersaline water bodies (>60)
(Grigg et al., 1986). Animals
maintain their osmotic balance by employing extrarenal lingual salt glands to
concentrate and remove excess salt accumulated as a consequence of living in
marine habitats. Data from the current and previous studies has shown that the
salt glands of C. porosus can respond rapidly to changes in
environmental salinity/plasma osmolarity
(Cramp et al., 2007;
Franklin and Grigg, 1993). The
salt glands of C. porosus respond to increasing environmental
salinity in much the same way as those of other marine reptiles and birds, in
that they become larger, have larger secretory cells, larger blood vessels
(Franklin and Grigg, 1993) and
show some evidence of neural plasticity upon hypertrophying
(Cramp et al., 2007). We have
shown that a change in the ultrastructure of salt gland secretory cells (i.e.
increased mitochondrial numbers and increase plasma membrane surface area)
correlates with an increased functional activity of the glands. Thus, the salt
glands of crocodiles exposed to hypersaline conditions have a greater
secretory rate and higher tissue metabolic rate than those of
freshwater-acclimated animals.
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Acknowledgments |
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References |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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