|
| |
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online February 15, 2006
Journal of Experimental Biology 209, 860-870 (2006)
Published by The Company of Biologists 2006
doi: 10.1242/jeb.02055
The accumulation of methylamine counteracting solutes in elasmobranchs with differing levels of urea: a comparison of marine and freshwater species
1 Ocean Sciences Centre, Memorial University of Newfoundland, St John's,
Newfoundland and Labrador, Canada A1C 5S7
2 Department of Integrative Biology, University of Guelph, Guelph, Ontario,
Canada N1G 2W1
3 Department of Cellular and Molecular Physiology, Yale University School of
Medicine, New Haven, Connecticut, USA 06520
4 Department of Biological Science, National University of Singapore, Kent
Ridge, Singapore 117543, Republic of Singapore
* Author for correspondence (e-mail: jtreberg{at}mun.ca)
Accepted 21 December 2005
 |
Summary |
|---|
 TOP Summary IntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
Key words: trimethylamine oxide (TMAO), betaine, trimethylamine oxidase, choline dehydrogenase, betaine aldehyde dehydrogenase, organic osmolyte, elasmobranch
|
Introduction |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
).
For example, as part of their osmoconforming osmoregulatory strategy, marine
elasmobranchs accumulate urea to concentrations as high as 300–500 mmol
l–1 (Yancey et al.,
1982; Yancey,
1994; Ballantyne,
1997). Urea can also reach levels greater than 1000 mmol
l–1 in the inner medullary region of the kidney of some
dehydration tolerant mammals (Yancey,
1988). It has been hypothesized that methylamine organic osmolytes
are accumulated in concert with urea to counteract the perturbing effects of
urea on protein structure (Yancey et al.,
1982; Yancey,
1994). Based on in vitro experiments, a urea to
`counteracting' solutes concentration ratio of about 2:1 may be optimal for
preserving proper protein function
(Yancey, 1994), but few data
on in vivo experimental tests of this ratio are available.
In a study of three rodent species
(Yancey, 1988) the urea
gradient found from the mammalian renal cortex to the inner medulla was
exploited as a test of the hypothesis that counteracting solutes protect
against urea-induced protein denaturation. Consistent with the hypothesis,
Yancey found that (i) the total methylamine content of the kidney increases
along the urea gradient, and (ii) there was a significant correlation between
total methylamine and urea content in the outer and inner medulla across
mammalian species (Yancey,
1988). However, the cortex of all three species had much lower
total methylamine content than would be expected based on the regression
analysis of the medullary regions, despite having significant urea content
(30–44 µmol g–1). Furthermore, the cortex and
medulla (outer and inner) are anatomically distinct tissues with different
functions and intracellular volumes, which complicates interpretations because
the accumulation of methylamines in the kidney is largely an intracellular
phenomenon (discussed by Yancey,
1988).
Similar to the mammalian kidney, the muscle of marine elasmobranchs also
has high non-urea organic osmolyte content. The methylamines trimethylamine
oxide (TMAO), betaine and sarcosine
(Withers et al., 1994;
Steele et al., 2005), as well
as ß-amino acids, taurine and ß-alanine, make up most of the
non-urea organic osmolyte component in marine elasmobranch muscle
(Yancey, 2001). As is the case
in the mammalian kidney, the muscle intracellular fluid of elasmobranchs
accumulates methylamines to a higher concentration than is found in the
extracellular fluid. It should be noted that methylamines are generally viewed
as counteracting solutes whereas ß-amino acids are considered compatible
solutes; that is, high concentrations of the latter have relatively little
effect on protein structure (Yancey,
1994). While the bulk of the non-urea organic osmolytes are
typically methylamines (see reviews by
Yancey et al., 1982;
Yancey, 1994;
Yancey, 2001), some marine
elasmobranch species require the inclusion of ß-amino acids to achieve or
approach the 2:1 ratio between urea and other solutes
(Steele et al., 2005).
Elasmobranchs are not strictly a marine group of fishes. Stingrays of the
family Potamotrygonidae, which have been isolated from their marine ancestors
in the Amazon basin for about 15–23 million years
(Lovejoy et al., 1998), have
entirely lost the physiological capacity to accumulate urea in their tissues
(Thorson et al., 1967;
Thorson, 1970) and are
considered stenohaline freshwater species, i.e. they are unable to survive in
salinities of more than about 40% seawater
(Thorson, 1970;
Tam et al., 2003). In contrast
to stenohaline elasmobranchs that are restricted to either marine or
freshwater environments, there are also euryhaline elasmobranchs capable of
living in both marine and freshwater habitats. Under marine conditions,
euryhaline elasmobranchs osmoregulate in a similar manner as strictly marine
species (Piermarini and Evans,
1998; Pillans et al.,
2005), but acclimation of a euryhaline species to freshwater is
associated with a marked decrease of body fluid osmolality that is largely
attributed to a reduction in the concentration of urea. Even once fully
adapted to freshwater, euryhaline elasmobranchs maintain levels of urea that
are quite high (approximately 140–200 mmol l–1)
(Thorson et al., 1973;
Piermarini and Evans, 1998:
Pillans and Franklin, 2004).
Recently, a third `intermediate' form of freshwater elasmobranch has been
discovered. Himantura signifer, also a stingray, is normally found in
freshwater but does venture into brackish water and still has the capacity for
urea accumulation (Tam et al.,
2003). Plasma urea in H. signifer in freshwater is about
45–70 mmol l–1. This species has higher salinity
tolerance than potamotrygonid stingrays, partially due to its ability to
elevate the osmolality of body fluids via further accumulation of
urea when acclimated to 60% seawater (Tam
et al., 2003; Ip et al.,
2005).
In parallel with a pattern of decreased capacity for urea accumulation in
freshwater elasmobranchs compared to marine species, there is a diminished
capacity for urea synthesis via the ornithine–urea cycle. For
example, the rate limiting step of urea synthesis in the liver,
carbamoylphosphate synthetase III (CPS III) activity, is very low in the
stenohaline freshwater Potamotrygon motoro compared to euryhaline and
marine species (Anderson, 1980;
Tam et al., 2003).
In contrast to urea, little is known about the synthesis of methylamines in
elasmobranchs. The ability to oxidize trimethylamine (TMA) to TMAO is
considered an essential component of TMAO synthesis, and the enzyme that
catalyzes this reaction is trimethylamine oxidase (TMAoxi), which occurs
sporadically in elasmobranchs (Baker et
al., 1963; Goldstein and
Dewitt-Harley, 1973). When present, TMAoxi activity is high in the
liver of elasmobranchs (Goldstein and
Dewitt-Harley, 1973), suggesting that this organ is a major site
of TMAO synthesis (Baker et al.,
1963; Goldstein and
Funkenhouser, 1972; Goldstein
and Dewitt-Harley, 1973). To our knowledge betaine synthesis has
never been studied in an elasmobranch. In mammals and other vertebrates,
betaine is synthesized from choline by the sequential and physiologically
irreversible dehydration reactions of choline dehydrogenase (ChoDH) and
betaine aldehyde dehydrogenase (BADH). Unlike the sporadic pattern of TMAoxi
activity, it is likely that the betaine synthetic pathway is found in most if
not all elasmobranchs, because it plays a critical role in the catabolism of
dietary choline (Haubrich and Gerber,
1981).
The physiological range of urea accumulation found in elasmobranchs, from several hundred millimolar in marine species to negligible amounts in the potamotrygonid stingrays, provides a unique opportunity to test the hypothesis that counteracting solutes are maintained in a conserved ratio with urea, while avoiding the complicating factors associated with the mammalian kidney (e.g. differing intracellular volumes). Therefore, the first goal of this study was to determine if specific methylamines, the total methylamine content, or the combined total of non-urea osmolytes (methylamines and the ß-amino acids taurine and ß-alanine) are maintained in a conserved ratio with urea in muscle tissue among elasmobranch species with varying degrees of intracellular urea accumulation.
The second goal of this study was to determine if the activity of enzymes
involved with methylamine synthesis (TMAO and betaine) in the liver vary in
elasmobranchs according to the degree that a particular methylamine is
accumulated in muscle, like the relationship between CPS III and urea (see
above). To accomplish this, we again exploited the interspecific variation in
methylamine accumulation across marine and freshwater elasmobranchs. We
focused on the synthesis of TMAO and betaine because these are the predominant
methylamines found in the muscle of elasmobranchs
(Withers et al., 1994;
Steele et al., 2005) and these
compounds are generally the most effective organic osmolytes for the
counteraction of urea effects on proteins
(Yancey, 1994). We
hypothesized that in species that lack TMAoxi activity, there may be elevated
betaine synthetic capacity. If methylamines are required to counteract urea
effects, the capacity for betaine synthesis may be important in species
lacking TMAoxi activity during periods of reduced dietary TMAO intake, in
order to supplement the muscle methylamine pool via betaine
synthesis.
|
Materials and methods |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
) tanks
maintained at 8–13°C for at least 3 months prior to sampling. During
this period, they were fed chopped herring 2–3 times per week to
satiation. Leucoraja erinacea Mitchill were caught by otter-trawl in
Passamaquoddy Bay, New Brunswick, Canada, transported to the Huntsman Marine
Science Laboratory and sampled within 5–7 days of capture. Animals were
housed in a flow-though seawater (approximately 31) tank
(11–14°C) without feeding.
Taeniura lymma Forsskål and Chiloscyllium punctatum
Müller and Henle were purchased at a live fish market in Singapore
approximately 6 h after capture from the ocean. They were transported to the
National University of Singapore (NUS), where they were held in full-strength
(30) seawater at 25°C in fibreglass aquaria under a 12 h:12 h
photoperiod. T. lymma were sampled within 3 days of acquisition;
C. punctatum were sampled the same day they were obtained. Water was
changed daily and the animals were not fed.
Euryhaline species in freshwater
Dasyatis sabina Lesueur were caught on baited trot-lines from the
St Johns River system, Florida, USA, and housed in freshwater tanks
(<1) as described (Piermarini
and Evans, 1998).
Freshwater species
Potamotrygon motoro Müller and Henle and Himantura
signifer Compagno and Roberts were obtained from a fish farm in
Singapore, where they were fed bloodworms (Chironomidae). They were
transported to NUS in water-filled plastic bags. Holding conditions were as
described (Tam et al., 2003).
Briefly, animals were held in freshwater at 25°C in plastic aquaria (3
animals per aquarium) under a 12 h:12 h photoperiod. Water was changed daily,
the animals were not fed. P. motoro were sampled within 3 days of
acquisition. Three days after specimen acquisition, a group of H.
signifer was gradually acclimated to half-strength seawater (15)
using the following regime: freshwater (day 1) to 5 (day 2) to
10 (day 3) to 15 (day 4, held until day 14). A control group
was kept in freshwater. Both the control and experimental groups comprised
animals in several separate aquaria to avoid tank effects. Animals were not
fed for the duration of the experiment. Animals from both groups were sampled
after 14 days of acclimation. All fish used in this study were in apparent
good condition and were maintained on a natural photoperiod unless otherwise
stated.
TMAO determination
Samples were homogenized in 9 volumes of ice-cold TCA (5%) with a motorized
tissue homogenizer and left on ice for 10 min to precipitate protein.
Following centrifugation, 15 600 g for 5 min, supernatants
were assayed directly for TMAO spectrophotometrically
(Wekell and Barnett, 1991), as
described previously (Sulikowski et al.,
2003; Treberg et al.,
2005). Elasmobranch tissues typically have low TMA content
compared to TMAO (Treberg and Driedzic,
2002), which was confirmed for muscle samples from L.
ocellata, L. erinacea and D. sabina (data not shown); thus
samples have not been corrected for endogenous TMA. For samples that were
expected to have very low TMAO content (P. motoro and H.
signifer), we further modified the assay to improve the lower limits of
detection. Tissues were prepared as described above, and 300 µl of TCA
extract was added to a 1.5 ml centrifuge tube followed by 300 µl of both
toluene and then the iron–EDTA reducing mixture as described
(Wekell and Barnett, 1991).
The tube was closed and heated at 50°C for 5 min. Tubes were allowed to
cool to room temperature, 600 µl of 45% KOH was added and the tube was
vortex mixed for 15 s. Tubes were vortexed twice more, after allowing
3–5 min between mixings for phase separation, to extract TMA into the
toluene phase. After the final mixing, 200 µl of the toluene phase was
added to another 1.5 ml centrifuge tube containing approximately 10–20
mg of sodium sulphate followed by 500 µl of 0.02% picric acid in toluene.
This was mixed and left for at least 2 min to remove any residual water and
the absorbance of 650 µl of the toluene phase was read at 410 nm.
Trimethylamine oxide content of tissues was determined based on the absorbance
at 410 nm compared to a standard curve (0–3 mmol l–1
for the standard assay and 0–0.3 mmol l–1 for the
higher resolution assay describe above).
HPLC analysis of organic osmolytes
The determination of all other organic osmolytes was done by HPLC using a
Waters SugarPak column and refractive index detector. Muscle samples were
homogenized in 9 volumes of 6% PCA and then prepared as described
(Wolff et al., 1989), where
the mobile phase is 50 mg l–1 Ca-EDTA. While this provided
suitable resolution of most osmolytes, we found that there was poor resolution
between betaine and alanine (Fig.
1). Tam et al. (Tam et al.,
2003) found that alanine content is significant (2 µmol
g–1) in the muscle of H. signifer. Since alanine
would potentially complicate betaine determinations in this species, as well
as P. motoro, we modified the method of Wolff et al.
(Wolff et al., 1989) to
improve the resolution of alanine and betaine.
|
Using a flow rate of 0.6 ml min–1, at a column temperature of 90°C, we tried a number of concentrations of Ca-EDTA in the mobile phase and found that a mobile phase of 500 mg ml–1 was suitable for the determination of all solutes in the present study (data not shown), except betaine and sarcosine because of little to no resolution from alanine and glycine, respectively (Fig. 1). However, a mobile phase of 25 mg l–1 Ca-EDTA allowed for suitable resolution of betaine, alanine, glycine and sarcosine (Fig. 1) if the column is fully equilibrated with calcium, which is a requirement to maintain resolution. By running a set of samples (4–6) at 500 mg l–1 followed by alternating to the 25 mg l–1 Ca-EDTA mobile phase and re-running the same samples, all of the major organic osmolytes (other than TMAO) could be assessed.
Enzyme activities
Liver samples were homogenized in 5 or 9 volumes of ice-cold buffer (50
mmol l–1 Hepes, 1 mmol l–1 EDTA, pH 7.5 at
20°C). For ChoDH and BADH, Triton X-100 was added to homogenates (0.2%
final concentration). Homogenates were used without centrifugation for ChoDH
and TMAoxi. For BADH, homogenates needed to be cleared by centrifuging (15 600
g for 10 min at 4°C) to reduce the light scattering
effects of cellular debris. All assays were conducted at 20°C and were
linear with time and protein.
Choline dehydrogenase, which catalyzes the conversion of choline to betaine
aldehyde, was assayed by a modification of the method used by Haubrich and
Gerber (Haubrich and Gerber,
1981). The assay mixture was composed of 50 mmol
l–1 phosphate buffer (pH 8.0), 2 mmol l–1
phenazine methosulphate (PMS) and 5 mmol l–1 choline,
including sufficient [14C-methyl]choline-HCl to give 0.125 µCi
ml–1 (1 Ci=3.7x1010 Bq) in the final assay
volume of 300 µl. The reaction was terminated by adding an equal volume
(300 µl) of ice-cold 6% PCA followed by centrifugation (15 600
g for 5 min). 500 µl of the supernatant was collected and
mixed with 45 µl of 6 mol l–1 KOH to remove most of the
perchlorate. This was centrifuged as above, the supernatant collected and
frozen (–20°C) for later analysis.
To determine the amount of [14C-methyl]betaine aldehyde produced, samples were thawed and duplicate 100 µl portions mixed with 20 µl of 1 mol l–1 NaOH and 20 µl of 30% hydrogen peroxide and incubated at room temperature for 60 min to chemically oxidize betaine aldehyde to betaine. To separate [14C-methyl]choline from [14C-methyl]betaine, a small ion-exchange column was made by adding 1 ml of a 1:1 slurry of BioRad AG-50W resin (Hercules, CA, USA; Li+ form, 200–400 mesh) and water to a Pasteur pipet with the tip blocked with a small piece of cotton. The column was packed with 2 ml of deionized water and the entire oxidized sample was added to the column. Labeled choline is retained by the resin while betaine was washed off with 2 ml of deionized water and collected into a 20 ml scintillation vial. The elutant was mixed with 10 ml of Ecolume (ICN, Irvine, CA, USA) followed by scintillation counting. The increase in radioactivity, relative to that found when PCA was added before initiating the reaction, was used as the measure of ChoDH activity. The formation of product was entirely dependent on the inclusion of the electron acceptor PMS (there was no difference in values obtained without PMS and when the reaction was immediately terminated after adding substrate).
A spectrophotometric assay (Perrino and
Pierce, 2000) was used to determine the BADH activity at 340 nm on
centrifuged homogenates. Control rates of absorbance change in the absence of
substrate were determined and subtracted in all cases.
A radioisotopic assay was developed, based on the assay of Baker and
Chaykin (Baker and Chaykin,
1962), to determine the TMAoxi activity of liver homogenates. The
assay conditions were as follows: 50 mmol l–1 Tris (pH 8.5),
50 mmol l–1 KCl, 0.3 mmol l–1 NADPH, 1.0
mmol l–1 TMA including [14C]TMA at a final
concentration of 0.1 µCi ml–1. Control values were
obtained by the omission of NADPH from the assay medium – product
formation is entirely dependent on NADPH (data not shown). The reaction was
stopped by combining the reaction mixture with an equal volume of ice-cold 5%
TCA. After 10 min, to allow complete precipitation of proteins, the mixture
was centrifuged at 15 600 g for 5 min at room temperature and
150 µl of the supernatant loaded onto a small column of BioRad AG-50W resin
(Na+ form, 200-400 mesh). Columns were made by adding 1 ml of a 1:1
slurry of resin and deionized water (0.5 ml bed volume) to a Pasteur pipet
with the tip blocked by a small amount of cotton. The resin was washed and
packed by two 1 ml volumes of deionized water prior to the addition of the TCA
extracted assay mixture. The TCA extract was loaded into the column and
non-exchangeable compounds removed by two 1 ml volumes of deionized water.
Radiolabelled TMA and TMAO were eluted with 2 ml of 0.1 mol
l–1 NaOH and the elutant collected into a 20 ml scintillation
vial. The elutant was heated to 70–80°C and evaporated to dryness in
a fumehood. Under alkaline conditions TMA is volatile whereas TMAO is not,
thus by evaporating the 0.1 mol l–1 NaOH elutant under
moderate heat the TMA is removed while the [14C]TMAO remains. The
residue was redissolved in 1 ml of deionized water and mixed with 10 ml of
scintillation cocktail (Ecolume) to determine the amount of TMAO produced by
scintillation counting.
The protein concentration of homogenates was determined using a commercial kit (BioRad Dc protein assay) using BSA as a standard.
Statistical analysis
When appropriate, means were compared by one-way analysis of variance
(ANOVA; P<0.05 was considered significant) and patterns across
species were assessed by linear regression.
|
Results |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
|
Euryhaline species in freshwater
The non-urea osmolytes in the muscle of D. sabina are
predominantly methylamines, mostly betaine, with ß-amino acids making
only a modest contribution.
Freshwater species
The most striking pattern in muscle of the two freshwater species examined
is that ß-amino acids are accumulated to a much greater extent than
methylamines, regardless of whether the species accumulates urea
(Fig. 2). Taurine is the
predominate ß-amino acid in both H. signifer and P.
motoro. Betaine was found at levels of 14.7±4.0 µmol
g–1 (mean ± s.e.m.) and 6.8±2.3 µmol
g–1 in the muscle of H. signifer acclimated to 50%
seawater and freshwater, respectively. Sarcosine was low in all freshwater
species (
1.0 µmol g–1 or less), as was TMAO:
0.67±0.15 µmol g–1 for H. signifer in 50%
seawater, 0.71±0.28 µmol g–1 H. signifer
in freshwater and 0.39±0.16 µmol g–1 in the muscle
of P. motoro.
Plasma methylamines in urea accumulating elasmobranchs in freshwater
Plasma concentrations of TMAO and betaine were 5.7±0.82 mmol
l–1 and 3.9±1.0 mmol l–1,
respectively, in D. sabina and were below the limits of detection in
H. signifer. This indicates that, like marine elasmobranchs (see
Withers et al., 1994), there
is a much lower methylamine concentration in the extracellular fluid than in
the intracellular fluid of the muscle in freshwater elasmobranchs.
Total methylamines and ß-amino acids in comparison to muscle urea content
Values for muscle urea content followed the expected decrease from marine
to freshwater species. Urea content ranged from 230 to 350 µmol
g–1 for marine species, was 146±7.0 µmol
g–1 in the euryhaline freshwater D. sabina, and was
96±6.5 and 75±3.1 µmol g–1 for 50% seawater
and freshwater acclimated H. signifer, respectively. Urea
accumulation in P. motoro was negligible (2.0±0.4 µmol
g–1) (Fig. 3).
No single methylamine or ß-amino acid correlated with urea across species
(data not shown). Total methylamines show a qualitatively similar trend across
species, as seen with urea, while total ß-amino acids do not. Although
total ß-amino acids appear to have a random distribution among marine and
euryhaline species, when a species has high total methylamines (for example
C. punctatum and D. sabina) it also has a proportionally
lower total ß-amino acids (Fig.
3).
|
The qualitative trend between muscle urea and total methylamine content (Fig. 3) is statistically supported (P<0.01) by linear regression analysis (Fig. 4), with a slope approaching that which would be expected for a 2:1 ratio between intracellular urea and total methylamines (see below). However, the fit of the data is modest (r2=0.65) compared to the regression of total methylamines plus total ß-amino acids (discussed below). Also, there are some species far off the overall trend (e.g. C. punctatum), and despite having much lower urea content, D. sabina has a total methylamine content similar to the marine skates and stingray. These observations do not support the notion of a highly conserved ratio between total methylamines and urea in elasmobranch muscle in all species. The data do illustrate that species with high levels of urea (i.e. marine species and D. sabina in freshwater), preferentially accumulate methylamines over ß-amino acids.
|
), would result in a slope of less than 0.5. Urea
concentration in the intracellular and extracellular fluid is similar and
would not be influenced in the same manner (see review by
Yancey, 2001).
Muscle organic osmolyte accumulation in H. signifer in response to salinity challenge
Although not the primary goal of this study, by examining the muscle
solutes in H. signifer that have been acclimated to 50% seawater in a
fasted state, it is possible to determine which solutes they can synthesize
and accumulate strictly from endogenous resources. When acclimated to 50%
seawater, muscle urea and taurine content significantly increased in H.
signifer. Mean betaine content more than doubled compared to freshwater
control animals (15±4.0 and 6.8±2.3 µmol
g–1, respectively). While this increase in betaine only
approached statistical significance (P=0.084), there are a number of
considerations that are suggestive of endogenous synthesis of a methylamine
solute in response to salinity challenge. This has not yet been demonstrated
in an elasmobranch in the absence of feeding. None of the other measured
methylamines increased in response to salinity challenge; however, the summed
total methylamines did significantly increase in H. signifer
acclimated to brackish water (Fig.
3). Betaine is the major constituent of the muscle methylamine
pool and the difference between total methylamines and betaine are very
similar, supporting the notion of betaine accumulation. Moreover, one of the
freshwater animals had unusually high muscle betaine content (22.2 µmol
g–1; based on the analysis of the other nine freshwater
specimens, this value is more than 3 s.d. higher than the mean) while all
other solutes for this individual were well within the `normal' range found in
this species. Exclusion of this individual does result in a significant
difference in betaine content between freshwater and 50% seawater groups.
Methylamine synthesizing enzyme activities
TMAO synthesis
Trimethylamine oxidase activity was below the limits of detection
(approximately 0.05 µmol h–1 g–1) for all
seven Rajiformes species (skates and stingrays, including members of the
families Rajidae, Dasyatidae and Potamotrygonidae) in the present study,
showing no trend between TMAO accumulation and synthetic capacity in these
taxa. The shark C. punctatum had high liver TMAoxi activity
(1.27±0.07 nmol min–1 mg protein–1 or
5.1± 0.87 µmol h–1 g–1) and also
had the highest level of TMAO in muscle
(Fig. 2 and
Table 1).
|
Betaine synthesis
All species examined had measurable liver ChoDH and BADH activity. Thus,
all species have a significant capacity for betaine synthesis. There is a
qualitative trend of increasing ChoDH and BADH activity with increasing
betaine content in those species able to tolerate full strength seawater
(marine and euryhaline species) but neither trend extends to the freshwater
species. Both freshwater species have high enzyme activity and relatively low
muscle betaine content (Fig.
5). Quantitatively, the relationship between liver ChoDH activity
and betaine accumulation across the species that can survive in full strength
seawater is not significant if analyzed on a species mean basis
(P>0.15, Fig. 5A)
or on data for individual animals (P=0.09; data not shown). The trend
with liver BADH activity and muscle betaine content approaches significance
using species means (P=0.08, Fig.
5B), and when analyzed on an individual basis is highly
significant across these five species (P<0.001,
r2=0.49; data not shown). Liver BADH activity appears to
be a better indicator than ChoDH of muscle betaine content in elasmobranchs
analyzed, and the above suggests that BADH activity is elevated to a greater
degree than ChoDH in betaine-accumulating elasmobranchs. When expressed as a
ratio of BADH/ChoDH, there is a significant relationship with muscle betaine
content on either a species mean or individual basis with all species
including the freshwater H. signifer and P. motoro
(Fig. 5C). Thus, across
species, as muscle betaine content increases, the enzymatic potential for
conversion of betaine aldehyde to betaine (BADH) significantly increases
relative to the capacity for the production of betaine aldehyde (ChoDH
activity). The only species with measurable TMAoxi activity, C.
punctatum, had low muscle betaine content and very low liver BADH
activity.
|
|
Discussion |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
). Consistent
with our observed shift from the accumulation of counteracting methylamines to
the compatible ß-amino acids at lower urea levels, Yancey and Burg
(Yancey and Burg, 1990) found
that the growth of cultured mammalian cells, both renal and non-renal, was not
inhibited by urea concentrations below 100–150 mmol
l–1. Despite the shift from methylamines to ß-amino acids as the major non-urea osmolytes, there is still a strong correlation between the total non-urea osmolytes (methylamines and ß-amino acids) and urea in the muscle of elasmobranchs (Fig. 4). This relationship is close to the expected 2:1 ratio when either the total methylamines or the combined values for methylamines and ß-amino acids are regressed with urea content. Overall, the combined total methylamines and ß-amino acids provides a substantially better fit than the total methylamines. We caution against overinterpreting the significance of the slope of methylamine regression because data for C. punctatum indicate that there is not a highly conserved ratio between urea and methylamines in the muscle of some marine elasmobranchs. That being said, it is clear that total methylamine content is higher in urea-accumulating elasmobranchs, even if there is not a tightly regulated ratio between methylamines and urea. Both of the above trends between urea and other organic osmolytes are consistent from marine species to the Potamotrygonid stingrays, with the intercept at negligible urea content being remarkably similar to the measured value for P. motoro (Fig. 4).
The distribution of TMAO synthesis in elasmobranchs
The lack of detectable TMAoxi activity in all of the skates and stingrays
examined in the present study is consistent with past surveys on TMAO
synthesis in elasmobranchs (Fig.
6 and references therein). The sporadic distribution of TMAoxi has
been known, but unexplained, for several decades
(Baker et al., 1963;
Goldstein and DeWitt-Harley,
1973). It was speculated by Goldstein and Palatt
(Goldstein and Palatt, 1974)
that `random deletions' may give an explanation for TMAoxi distribution, and
they drew an analogous comparison with ascorbic acid biosynthesis in mammals
where some higher taxa have lost the capacity for physiologically sufficient
endogenous synthesis. Fig. 6
illustrates how the available data on the presence or absence of detectable
capacity for TMA oxidation (in vitro or in vivo) is directly
linked to phylogeny in chondrichthyans (elasmobranchs and the chimaeras or
holocephalans). While further species need to be studied to confirm the
distribution postulated in Fig.
6, as well as to fill in `gaps' such as missing orders and
families, we propose that the distribution of TMAoxi in elasmobranchs can be
explained when grouped taxonomically.
|
Capacity for betaine synthesis and muscle accumulation
The present data indicate that an increase in the activity of liver BADH,
relative to ChoDH, coincides with the accumulation of betaine in the muscle of
elasmobranchs (Fig. 5C),
suggesting that BADH plays a regulatory role in betaine synthesis. The uptake
of choline by rat liver mitochondria, and thus the exogenous supply of
choline, is believed to be the major control site of betaine synthesis (Kaplan
et al., 1993). However, rat liver cells do not synthesize betaine in response
to hyperosmotic challenge (Wettstein et
al., 1998). Rabbit kidney cells (TALH) do synthesize and
accumulate betaine when exposed to hyperosmotic conditions
(Grunewald and Eckstein, 1995)
and are found in a `high urea' environment, making them a better analogy to
elasmobranch betaine synthesis. Kidney cells under high osmolality conditions
exhibit little or no activation of ChoDH, whereas BADH activity increases to 3
times that found in cells in isoosmotic (300 mOsm) medium. Grunewald and
Eckstein (Grunewald and Eckstein,
1995) surmized that BADH may be the rate limiting enzyme of
betaine synthesis in mammalian kidney cells under these conditions. Our
observation that BADH activity relative to ChoDH (the initial enzymatic step
of betaine synthesis) increases with betaine accumulation in elasmobranchs is
consistent with the notion that BADH activity may be rate limiting in
elasmobranchs; however, the supply of choline to the liver as a precursor to
betaine may still be of critical importance in the control of actual in
vivo betaine synthesis.
Curiously, there was no change in the activities of ChoDH or BADH in H. signifer acclimated to 50% seawater compared to freshwater acclimated animals. If activation of liver BADH is indicative of betaine synthesis in marine elasmobranchs, it is unclear why this would not also be the case in a freshwater species. It is possible that the high constitutive enzyme activities in this species preclude any need for activation in order to facilitate flux from choline to betaine. Furthermore, the liver supply of choline precursors and endogenous synthesis of choline may provide the requisite materials for any increased synthesis of betaine.
Although the liver is the major site of betaine synthesis in mammals
(Haubrich and Gerber, 1981),
it is important to note the possibility that other tissues not included in the
present study, or alternate metabolic pathways, may be involved with the
synthesis of methylamines in elasmobranchs. That stated, substantial
constitutive capacity for betaine synthesis via choline degradation,
and thus potentially sarcosine synthesis as well
(King et al., 1980), was found
in all elasmobranchs examined.
Exogenous or endogenous origins for methylamines?
The relative importance of endogenous methylamine synthesis to their
accumulation in elasmobranchs is still in need of clarification, but our data
do offer some preliminary insights on the source of methylamines. The only
species in this study that was fully acclimatized to laboratory conditions for
several months was L. ocellata. Specimens of L. ocellata
were fed chopped herring, which is a rich source of TMAO (approximately
25–40 µmol g–1) that has very low levels (<1
µmol g–1) of betaine and sarcosine
(Carr et al., 1996). When
compared to the congeneric L. erinacea, which were sampled shortly
after capture and fed on a natural diet of invertebrates that are high in
betaine (Carr et al., 1996;
Collette et al., 2004), specimens of L. ocellata were characterized
by higher TMAO and lower betaine and sarcosine levels. No skates examined to
date have exhibited demonstrable capacity for the conversion of TMA to TMAO,
yet both species have betaine and likely sarcosine synthetic capacity
(Table 1;
King et al., 1980). It is
tempting to interpret these differences between congeneric skates as due to a
coupling between dietary derived and endogenously synthesized methylamine
accumulation in the muscle of skates. In this scenario, when insufficient
amounts of TMAO, or betaine, can be obtained from the diet, the remaining
required methylamine solutes are made up by endogenously synthesized betaine
and sarcosine; albeit the synthesis of these solutes may require exogenous
choline or other precursory metabolites
(King et al., 1980). The
muscle of D. sabina had modest levels of TMAO and high betaine
content. This species apparently lacks the capacity for TMAO synthesis, but
can synthesize betaine like the skates above
(Table 1). Most freshwater fish
and invertebrates have low levels of TMAO (reviewed by
Hebard et al., 1982), thus
these euryhaline stingrays caught in freshwater likely have a diet that is low
in TMAO. As with the above skates, it is reasonable to suggest that these
stingrays compensate for any `methylamine-gap' in the muscle by accumulating
endogenously synthesized betaine. The above postulations can also be extended
and simplified in the case of C. punctatum, where the high muscle
TMAO content could be reflective of the capacity for endogenous TMAO synthesis
and thus maintenance of high levels irrespective of dietary TMAO content.
However, while the above scenarios may be logically reasonable, it is
important to acknowledge that at present they are still hypothetical and need
to be experimentally tested.
|
List of abbreviations |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
|
Acknowledgments |
|---|
|
References |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionList of abbreviationsReferences |
|---|
Anderson, P. M. (1980). Glutamine- and
N-acetylglutamate-dependent carbamoyl phosphate synthetase in elasmobranchs.
Science 208,291
-293.
Baker, J. R. and Chaykin, S. (1962). The
biosynthesis of trimethylamine-N-oxide. J. Biol. Chem.
237,1309
-1313.
Baker, J. R., Struempler, A. and Chaykin, S. (1963). A comparative study of trimethylamine-N-oxide biosynthesis. Biochim. Biophys. Acta 71, 58-64.[Medline]
Ballantyne, J. S. (1997). Jaws: the inside story. The metabolism of elasmobranch fishes. Comp. Biochem. Physiol. 118B,703 -742.[CrossRef]
Carr, W. E. S., Netherton, J. C., III, Gleeson, R. A. and Derby, C. D. (1996). Stimulants of feeding behaviour in fish: analysis of tissues of diverse marine organisms. Biol. Bull. 190,149 -160.[Abstract]
Collette, B. and Klein-MacPhee, G. (2002). Bigelow and Schroeder's fishes of the Gulf of Maine (3rd edn). Washington DC: Smithsonian Institution Press.
Forster, R. P. and Goldstein, L. (1976).
Intracellular osmoregulatory role of amino acids and urea in marine
elasmobranchs. Am. J. Physiol.
230,925
-931.
Goldstein, L. and Funkenhouser, D. (1972). Biosynthesis of trimethylamine oxide in the nurse shark, Ginglymostoma cirratum. Comp. Biochem. Physiol. 42A, 51-57.
Goldstein, L. and Dewitt-Harley, S. (1973). Trimethylamine oxidase of nurse shark liver and its relationship to mixed function amine oxidase. Comp. Biochem. Physiol. 45B,895 -903.[CrossRef][Medline]
Goldstein, L. and Palatt, P. J. (1974).
Trimethylamine oxide excretion rates in elasmobranchs. Am. J.
Physiol. 227,1268
-1272.
Goldstein, L., Hartman, S. C. and Forster, R. P. (1967). On the origins of trimethylamine oxide in the spiny dogfish, Squalus acanthias. Comp. Biochem. Physiol. 21,719 -722.[CrossRef]
Grunewald, R. W. and Eckstein, A. (1995). Osmotic regulation of the betaine metabolism in immortalized renal cells. Kidney Int. 48,1714 -1720.[Medline]
Haubrich, D. R. and Gerber, N. H. (1981). Choline dehydrogenase. Assay, properties and inhibitors. Biochem. Pharm. 30,2993 -3000.[CrossRef][Medline]
Hebard, C. E., Flick, G. J. and Martin, R. E. (1982). Occurrence and significance of trimethylamine oxide and its derivatives in fish and shellfish. In Chemistry and Biochemistry of Marine Food Products (ed. R. E. Martin), pp.149 -175. Westport: AVI Publishing.
Ip, Y. K., Tam, W. L., Wong, W. P. and Chew, S. F. (2005). Marine (Taeniura lymma) and freshwater (Himantura signifer) elasmobranchs synthesize urea for osmotic water retention. Physiol. Biochem. Zool. 78,610 -619.[CrossRef][Medline]
Kaplin, C. P., Porter, R. K. and Brand, M. D. (1993). The choline transporter is the major site of control of choline oxidation in isolated rat liver mitochondria. FEBS Lett. 321,24 -26.[CrossRef][Medline]
King, P. A., Cha, C. and Goldstein, L. (1980). Amino acid metabolism and cell volume regulation in the little skate, Raja erinacea. J. Exp. Zool. 212, 79-86.
Lovejoy, N. R., Bermingham, E. and Martin, A. P. (1998). Marine incursion into South America. Nature 396,421 -422.[CrossRef]
Perrino, L. A. and Pierce, S. K. (2000). Betaine aldehyde dehydrogenase kinetics partially account for oyster population differences in glycine betaine synthesis. J. Exp. Zool. 286,238 -249.[CrossRef][Medline]
Piermarini, P. M. and Evans, D. H. (1998). Osmoregulation of the Atlantic Stingray (Dasyatis sabina) from the freshwater Lake Jesup of the St Johns River, Florida. Physiol. Zool. 71,553 -560.[Medline]
Pillans, R. D. and Franklin, C. E. (2004). Plasma osmolyte concentrations and rectal gland mass of bull sharks Carcharhinus leucas, captured along a salinity gradient. Comp. Biochem. Physiol. 138A,363 -371.[CrossRef]
Pillans, R. D., Good, J. P., Anderson, W. G., Hazon, N. and Franklin, C. E. (2005). Freshwater to seawater acclimation of juvenile bull sharks (Carcharhinus leucas): plasma osmolytes and Na+/K+-ATPase activity in gill, rectal gland, kidney and intestine. J. Comp. Physiol. 175B,37 -44.
Steele, S. L., Yancey, P. H. and Wright, P. A. (2005). The little skate Raja erinacea exhibits an extrahepatic ornithine urea cycle in the muscle and modulates nitrogen metabolism during low-salinity challenge. Physiol. Biochem. Zool. 78,216 -226.[CrossRef][Medline]
Sulikowski, J. A., Treberg, J. R. and Howell, W. H. (2003). Fluid regulation and physiological adjustments in the winter skate, Leucoraja ocellata, following exposure to reduced environmental salinities. Environ. Biol. Fish. 66,339 -348.[CrossRef]
Tam, W. L., Wong, W. P., Loong, A. M., Hiong, K. C., Chew, S.
F., Ballantyne, J. S. and Ip, Y. K. (2003). The
osmotic response of the Asian freshwater stingray (Himantura
signifer) to increased salinity: a comparison with marine (Taeniura
lymma) and Amazonian freshwater (Potamotrygon motoro) stingrays.
J. Exp. Biol. 206,2931
-2940.
Thorson, T. B. (1970). Freshwater stingrays, Potamotrygon spp.: failure to concentrate urea when exposed to saline medium. Life Sci. 9,893 -900.[CrossRef]
Thorson, T. B., Cowan, C. M. and Watson, D. E.
(1967). Potamotrygon spp.: elasmobranchs with low urea
content. Science 158,375
-377.
Thorson, T. B., Cowan, C. M. and Watson, D. E. (1973). Body fluid solutes of juveniles and adults of the euryhaline bull shark Carcharhinus leucas from freshwater and saline environments. Physiol. Zool. 46, 29-42.
Treberg, J. R. and Driedzic, W. R. (2002). Elevated levels of trimethylamine oxide in deep-sea fish: evidence for synthesis and intertissue physiological importance. J. Exp. Zool. 293,39 -45.[CrossRef][Medline]
Treberg, J. R., Bystrianski, J. S. and Driedzic, W. R. (2005). Temperature effects on trimethylamine oxide accumulation and the relationship between plasma concentration and tissue levels in smelt (Osmerus mordax). J. Exp. Zool. 303,283 -293.[CrossRef]
Wekell, J. C. and Barnett, H. (1991). New method for analysis of trimethylamine oxide using ferrous sulphate and EDTA. J. Food Sci. 56,132 -138.[CrossRef]
Wettstein, M., Weik, C., Holneicher, C. and Häussinger, D. (1998). Betaine as an osmolyte in rat liver: metabolism and cell-to-cell interactions. Hepatology 27,787 -793.[CrossRef][Medline]
Winchell, C. J., Martin, A. P. and Mallatt, J. (2004). Phylogeny of elasmobranchs based on LSU and SSU ribosomal RNA genes. Mol. Phylogenet. Evol. 31,214 -224.[CrossRef][Medline]
Withers, P. C., Morrison, G., Hefter, G. T. and Pang, T. (1994). Role of urea and methylamines in buoyancy of elasmobranchs. J. Exp. Biol. 188,175 -189.[Abstract]
Wolff, S., Yancey, P. H., Stanton, T. S. and Balaban, R. (1989). A simple HPLC method for quantitating the major organic solutes of the renal medulla. Am. J. Physiol. 256,F954 -F956.
Yancey, P. H. (1988). Osmotic effectors in kidneys of xeric and mesic rodents: corticomedullary distributions and changes with water availability. J. Comp. Physiol. B 158,369 -380.[CrossRef][Medline]
Yancey, P. H. (1994). Compatible and counteracting solutes. In Cellular and Molecular Physiology of Cell Volume Regulation (ed. K. Strange), pp.81 -109. Boca Raton: CRC Press.
Yancey, P. H. (2001). Nitrogen compounds as osmolytes. In Fish Physiology, Vol.20 , Nitrogen Excretion (ed. P. Wright and P. Anderson), pp. 309-341. San Diego: Academic Press.
Yancey, P. H. and Burg, M. B. (1990). Counteracting effects of urea and betaine in mammalian cells in culture. Am. J. Physiol. 258,R198 -R204.
Yancey, P. H., Clark, M. E., Hand, S. C., Bowlus, R. D. and Somero, G. N. (1982). Living with water stress: evolution of osmolyte systems. Science 217,1212 -1222.
Related articles in JEB:
This article has been cited by other articles:
|
|
 |
|
  N. J. Serkova, J. C. Rose, L. E. Epperson, H. V. Carey, and S. L. Martin Quantitative analysis of liver metabolites in three stages of the circannual hibernation cycle in 13-lined ground squirrels by NMR Physiol Genomics, September 11, 2007; 31(1): 15 - 24. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. R. Treberg and W. R. Driedzic Maintenance and accumulation of trimethylamine oxide by winter skate (Leucoraja ocellata): reliance on low whole animal losses rather than synthesis Am J Physiol Regulatory Integrative Comp Physiol, December 1, 2006; 291(6): R1790 - R1798. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. Speers-Roesch, Y. K. Ip, and J. S. Ballantyne Metabolic organization of freshwater, euryhaline, and marine elasmobranchs: implications for the evolution of energy metabolism in sharks and rays J. Exp. Biol., July 1, 2006; 209(13): 2495 - 2508. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Phillips ELASMOBRANCHS BALANCE UREA J. Exp. Biol., March 1, 2006; 209(5): ii - ii. [Full Text] [PDF]
|
|
| ||||||||||||||||||||
TMAO synthesis was assessed by in vivo
conversion of labeled precursor; –, below limits of detection or
questionable synthetic capacity; +, significant TMAO synthesis present. Values
in parentheses indicate the source of data:
1Baker et al., 1963