General and Comparative Endocrinology 153 (2007) 392–400
www.elsevier.com/locate/ygcen
Minireview
Measurement of fish steroids in water—a review
Alexander P. Scott *, Tim Ellis
The Centre for Environment, Fisheries and Aquaculture Science, Weymouth, Dorset, DT4 8UB, UK
Received 13 September 2006; revised 7 November 2006; accepted 13 November 2006
Available online 22 December 2006
Abstract
Measurement of fish steroids in water provides a non-invasive alternative to measurement in blood samples, offering the following
advantages: zero or minimal intervention (i.e. no anaesthetic, bleeding or handling stress); results not being biased by sampling stress;
repeat measurements on the same fish; the possibility of making non-lethal measurements on small and/or rare fish; integrating the
response of many (or of single) fish; and allowing concurrent monitoring of behaviour or physiology. The procedure is relatively new
and, although applications are still fairly limited, there are several themes and potential problem areas that are worthy of review.
Crown Copyright Ó 2006 Published by Elsevier Inc. All rights reserved.
Keywords: Fish; Steroids; Water; Non-invasive; Cortisol; 11-Ketotestosterone
1. History of the procedure
The impetus for the measurement of steroids in water
came from research on steroid pheromones. Stacey and
co-workers had proposed that the teleost oocyte maturation-inducing steroid, 17,20b-dihydroxypregn-4-en-3-one
(17,20b-P) was a male priming pheromone in goldfish (Carassius carassius) and wanted to prove that it was indeed
released into the water by females during the periovulatory
period. They established that this steroid was released in
relatively large amounts by ovulating females and that
the pattern of release of the steroid matched its pattern
of secretion into the plasma (Dulka et al., 1987; Stacey
et al., 1989). They also showed that periovulatory goldfish
released two other potential steroidal pheromones, 17-hydroxyprogesterone and 17,20b-P glucuronide (Van Der
Kraak et al., 1989). Working on the same species, Scott
and Sorensen showed that males and females released not
only these, but a wide range of other steroids in free, glucuronidated and sulphated forms (Scott and Sorensen, 1994;
Sorensen et al., 2005; Sorensen and Scott, 1994).
*
Corresponding author. Fax: +44 1305 206601.
E-mail address: [email protected] (A.P. Scott).
Still on the theme of pheromones, papers have also been
published on the release of steroids into the water by the
male peacock blenny (Salaria pavo) (Oliveira et al., 1999),
the tench (Tinca tinca) (Pinillos et al., 2003) and the roach
(Rutilus rutilus) (Lower et al., 2004). The last two studies
showed that, although the same types of steroids are found
in both cyprinid species, their rates and patterns of release
differ greatly from each other and from the goldfish. For
example, in male goldfish, very little 17,20b-P is released
into the water at any stage of reproduction. In roach, however, it is released in relative abundance by both sexes and
does not appear to be associated solely with the spawning
period (as it is with female goldfish). The male goldfish also
releases androstenedione (Ad) in amounts that far exceed
that of males of the other two species (Sorensen et al.,
2005).
In the course of the above-mentioned studies, it became
apparent (Oliveira et al., 1999; Oliveira et al., 2001; Scott
et al., 2001) that it did not matter if the steroids were pheromones or not, their measurement in water could be a useful way of studying reproductive physiology without
having to bleed the fish. It was shown that concentrations
of 17,20b-P in water correlated well with concentrations of
17,20b-P in plasma of individual female dentex (Dentex
dentex) that had been injected with gonadotropin-releasing
0016-6480/$ - see front matter Crown Copyright Ó 2006 Published by Elsevier Inc. All rights reserved.
doi:10.1016/j.ygcen.2006.11.006
A.P. Scott, T. Ellis / General and Comparative Endocrinology 153 (2007) 392–400
hormone (GnRH) (Greenwood et al., 2001) and could thus
be used to monitor the effectiveness of such treatments.
Water steroid measurements were later used to investigate
the timing of ovulation in populations of dentex that were
held at different sex ratios (Pavlidis et al., 2004) and to confirm the state of maturity of males and females in the European eel (Anguilla anguilla) (Huertas et al., 2006). In a
variety of cichlid species (Bender et al., 2006; Hirschenhauser et al., 2004; Oliveira et al., 2001, 2003), the Siamese
fighting fish (Betta splendens) (Dzieweczynski et al., 2006)
and bluebanded goby (Lythrypnus dalli) (Black et al.,
2005; Rodgers et al., 2006) measurements of 11-ketotestosterone (11-KT), and in most cases also testosterone (T), in
water have been used to investigate behavioural
endocrinology.
One of the steroids that was found to be released into the
water by goldfish was cortisol (Sorensen and Scott, 1994). Its
discovery in water in microgram quantities seeded the idea
for the non-invasive stress assay in fish based upon measurement of free cortisol in water—first proposed in 2001 (Scott
et al., 2001) and validated for rainbow trout in 2004 (Ellis
et al., 2004). Research had already shown that fish excrete
metabolised steroids into the water via the urine and bile
(Pottinger et al., 1992; Scott and Liley, 1994; Scott and Vermeirssen, 1994; Oliveira et al., 1996; Vermeirssen and Scott,
1996) and measurement of cortisol metabolites in faeces had
been tried, but with limited success, in fish (Oliveira et al.,
1999; Turner et al., 2003). As in the case of the sex steroids
(see above), it was confirmed that free cortisol is primarily
released into the water via the gills (Ellis et al., 2005).
The non-invasive stress assay has so far been applied to
stocking density experiments on carp (Ruane and Komen,
2003), to the effects of ship noise on European freshwater
fishes (Wysocki et al., 2006), to the effects of tag attachments in roach and carp (Lower et al., 2005) and to behavioural experiments on two cichlids, Archocentrus
nigrofasciatus (Earley et al., 2006) and Neolamprologus pulcher (Bender et al., 2006). It has also been extensively
applied to stocking density and disease challenge
experiments in our own laboratory, and these data will
be published in due course.
Using existing immunoassay procedures, concentrations
of cortisol and other steroids in water are, in all but
extreme situations, too low to be measured directly. This
means that steroids need to be extracted and concentrated
from the water. At the moment, this is time-consuming and
requires relatively expensive single-use solid-phase extraction cartridges and pre-filters, and the use of organic solvents (Ellis et al., 2004). Obviously, this limits the use of
the procedure to well-equipped scientific laboratories. This
fact has been used as an indictment of the procedure as a
putative operational indicator of fish welfare on fish farms
(Huntingford et al., 2006). However, this may be a shortsighted view. There are no theoretical barriers to the eventual development of a procedure for measuring cortisol (or
indeed any other compound that fish release into water)
that involves no extraction and no technical equipment.
393
Such a procedure (in the form of ‘sticks’ or ‘colour strips’
that are dipped into urine) is used every day all over the
world to test for pregnancy. The technical challenge in
developing a rapid test for cortisol in water will undoubtedly be greater than for human chorionic gonadotrophin in
urine. However, if the procedure answers a need for a diagnostic test for fish ‘well-being’, then the challenge can
undoubtedly be met. Dipstick tests may also be of practical
value in assessing the maturity status of broodstock fish
prior to handling and stripping (Huertas et al., 2006).
2. Measurement of free vs conjugated steroids
In several recent studies (Bender et al., 2006; Black et al.,
2005; Dzieweczynski et al., 2006; Hirschenhauser et al.,
2004; Oliveira et al., 2003) measurements were made of
free, sulphated and glucuronidated steroids and the concentrations were combined for reporting. We question the
need to measure the conjugated steroid fraction in water.
Vermeirssen and Scott (1996) showed that when rainbow
trout were injected with either a free, sulphated or glucuronidated steroid, the free steroid appeared in the water
via the gills, the sulphated steroid appeared in the urine
and the glucuronidated steroid accumulated in the bile.
By ‘bisecting’ goldfish and rainbow trout in specially constructed tanks, it was shown (Ellis et al., 2005; Sorensen
et al., 2000; Vermeirssen and Scott, 1996) that free steroids
were released only from the anterior (gill) region and sulphated and glucuronidated steroids only from the posterior
region. These findings suggest that free steroids in the water
represent passive ‘leakage’ of steroid across the gills as a
result of a concentration gradient between plasma and
water. On this basis, we argue that the concentration of
free steroid in the water equates to the concentration of
‘physiologically active’ steroid in the plasma very close to
the moment in time that the sample is taken. On the other
hand, concentrations of glucuronidated and sulphated steroid will not only be strongly influenced by the rates of urination and defaecation of the fish, but also by the time lag
required for the conjugation reaction to occur. For example, a delay of ca. 3 h has been demonstrated between
peaks of free and conjugated 17,20b-P excretion in periovulatory goldfish (Scott and Sorensen, 1994; Stacey et al.,
1989).
3. Advantages of measuring free steroids in water
Measurement of steroids in water, rather than in blood
samples taken from fish, offers the advantages of zero or
minimal intervention (i.e. no anaesthetic, bleeding or handling stress), the results not being biased by sampling
stress, repeat measurements on the same fish, the possibility
of making non-lethal measurements on small and/or rare
fish, the ability to integrate the response of many fish and
allowing concurrent behavioural or physiological monitoring. Measurement of steroids in water also comprises integration over time, thereby reducing the problem of short
394
A.P. Scott, T. Ellis / General and Comparative Endocrinology 153 (2007) 392–400
term (minute timescale) fluctuations in hormone levels that
may occur in the plasma (Dulka et al., 1987; Bender et al.,
2006). In terms of practical applications, as mentioned
above, a rapid test for cortisol in water has potential as
an operational indicator of fish welfare.
A recent development that may provide another means
for measuring steroids in water involves passive samplers
that are designed to be placed in situ and adsorb polar
organic chemicals over long periods of time (Alvarez
et al., 2004). Rather than processing a water sample taken
at a single time point, such integrative samplers have the
potential to provide an integrated ‘‘hormonal history’’ of
a fish population over a period of days-weeks; however,
we are not aware of their application in the types of studies
covered by this review. Their main use to date has been for
the detection of xenoestrogens in the environment (e.g.
Vermeirssen et al., 2005).
4. Potential problems in measuring and interpreting steroid
concentrations in water
The release of steroids is likely to be influenced not only
by the concentration in the plasma, but also to some degree
by their affinity for specific steroid binding proteins in the
plasma, the rate of blood flow through the gills, opercular
beat rate, fish size/gill surface area, gill permeability, gill
damage, salinity, water temperature and concentration gradient. There are little or no published data on any of these,
apart from the effect of sex steroid binding proteins on
release/uptake of steroids (Scott et al., 2005). The concentration of steroid in the water to be measured will be affected by all the above—and also by fish biomass, water
volume, water flow rate (in flow through systems), steroid
re-uptake by the fish (in static systems), steroid degradation
and steroid adsorption to surfaces. Measured concentrations could also be affected by instability of the steroid during storage and/or extraction and also by the specificity
and sensitivity of the assay. Most problems can be negated
by designing experiments so that tank volumes, water flow
rates, fish size, fish biomass and temperature are identical.
Certain problems can be solved in ways that are discussed
below.
Other potential problems with the water extraction
procedure that are not further discussed are: it is (presently) more costly and time-consuming than extraction
from plasma; it requires a lot more validation—an additional cost and workload; estimating steroid release rates
from water steroid values in a dynamic situation requires
calculation steps that distance the final data set from the
immunoassay results; measurement of steroids in water
effectively integrates the response of all fish within a tank
and thus information on individual variation is lost and
outlier individuals with high release rates could potentially bias results. These potential problems need to balanced against the advantages of measuring steroids
from water when deciding suitability for a particular
application.
5. The concentration of steroid in the water is dependent not
just on the physiological state of fish but on fish mass and
water flow rate
Water steroid concentration will depend not only upon
the release rate by the fish, but also on the fish biomass
and the dilution dynamics (tank volume and water replacement). Water steroid concentration (e.g. ng/L) is therefore
a relative measure that can only be used to make comparisons over time or in tanks with the same biomass, volume
and inflow rate (i.e. under carefully controlled conditions).
Although this can be achieved by good experimental
design, it is preferable to obtain an absolute measure of
the rate of steroid release (ng/g/h) to enable comparisons
between studies. Two ways in which the rate of steroid
release can be calculated are commonly used, and will
depend upon the system and fish biomass.
The simplest way of measuring steroid release rate is
to move a fish into a static volume of clean water for
a known period of time and then return it to its tank.
The steroid can then be extracted and assayed. This procedure has already been used for behavioural studies in
small fish where concentrations within the larger holding
tank would not be practical to measure (Dzieweczynski
et al., 2006; Hirschenhauser et al., 2004; Oliveira et al.,
2003; Rodgers et al., 2006). When used to measure 11KT and T, there would appear to be few problems with
this procedure, as the release of these steroids is unlikely
to be affected in the short-term (up to 30–60 min) by
handling and ‘confinement’. However, when used to measure cortisol (Bender et al., 2006; Rodgers et al., 2006)
that is responsive to stress, its application is contentious—as the act of transferring and subsequently confining the fish in a new environment would itself be
expected to affect cortisol concentrations. Ideally cortisol
status should be assessed from the undisturbed holding
tank.
In tanks with a higher fish biomass, water hormone concentration can be measured directly, although in a closed
system interpretation of values is complicated by the lack
of knowledge on the dynamics of removal (i.e. rate of
breakdown). The rate of steroid release can however be calculated in a dynamic situation (i.e. when water is flowing
through the tank). If the concentration is assumed not to
change over time, then release rate can be calculated
simply:
Release rateðe:g: pg=hÞ ¼ C R;
where C, concentration (pg/L); and F, flow rate (L/h).
However, if steroid concentrations are changing, at least
two measurements of water steroid concentration made at
a defined time interval are needed, the system volume must
be included in the calculation, and a more sophisticated
equation is required:
Release rate ¼ ðVktðC t C 0 ekt ÞÞ=ð1 ekt Þ
A.P. Scott, T. Ellis / General and Comparative Endocrinology 153 (2007) 392–400
where V is the water volume, C0 and Ct are the hormone
concentrations at the beginning and end of the sampling
period (over a time interval t) and k is the instantaneous
rate of loss due to dilution from the inflow water. The value
of k is F/V (for derivation, plus experimental validation,
see Ellis et al., 2004).
Obviously, values for total release rate calculated
through the above equations need to be corrected for the
biomass of fish, to express release rate in terms of amount
of steroid released per unit weight of fish per unit of time
(usually ng/g/h or pg/g/h).
In many situations (e.g. on commercial fish farms) it is
impossible to obtain accurate information on fish biomass
and water flow rate. A potential solution to this problem is
to use some form of ‘internal standard’ (‘housekeeper’ or
‘normaliser’) compound that is also released into the water
by fish, that is related to fish biomass, but is unaffected by
factors such as age, gender, feeding, reproduction or stress.
Steroid concentrations can then be expressed as a ratio to
the concentrations of this ‘housekeeping’ compound. Turner et al. (2003) attempted to apply this concept to correct
the cortisol content of parrotfish faeces by expressing it
as a ratio to T. Analogous problems have been faced by
researchers in a number of other biological fields. In molecular biology, for example, the expression of specific genes is
often normalised against b-actin. A normaliser for cortisol,
if one can be found, should hopefully also provide an
intrinsic correction for environmental factors, such as temperature, that are likely to affect the rate of release of steroids. One potential candidate that we have been
investigating is melatonin—a hormone synthesised by the
pineal gland and released in response to darkness. We have
already shown that melatonin is released into the water by
rainbow trout, that the amounts that are released during
the night are higher than those released during the day
and are stable (James et al., 2004). Melatonin is also
released into the water via the gills (Ellis et al., 2005)—
i.e. by the same mechanism as free steroids (Sorensen
et al., 2000; Vermeirssen and Scott, 1996). The results are
promising but the research is still very much ‘in progress’.
One drawback of using melatonin is that water samples
have to be collected during the night when the fish are
actually releasing melatonin.
6. Instability of steroids during extraction
Our research on the development of a non-invasive procedure for the assessment of fish welfare (Ellis et al., 2004)
was held up for well over a year by problems with the
recovery of cortisol from water. The fact that the problem
was intermittent made it very difficult to track down. Eventually, we discovered that it was the diethyl ether that we
were using to elute the solid-phase extraction cartridges.
We had chosen this solvent because it would only elute free
steroids from the cartridges (thus avoiding potential problems with cross-reacting conjugated steroids that might be
in the water) and also would evaporate rapidly in the fume
395
cupboard. By using tritiated cortisol, we showed that the
problem was not with the efficiency of extraction of cortisol
from the cartridges. The problem was that the immuno-reactivity of the (non-tritiated) cortisol being eluted was
being affected in some way that prevented its measurement.
We noted that the problem of intermittency was more frequent when we used ultrapure diethyl ether stabilised with
copper foil and hypothesised that the problem was due to
the formation of damaging peroxide radicals once the
diethyl ether had been transferred to the dispenser (in
which there was no copper foil). Although the problem
seemed to disappear when we used analytical-grade diethyl
ether stabilised with ethanol, we decided to opt on the side
of caution and switch to ethyl acetate instead. There are
three cautionary notes arising from this experience. Firstly,
‘high purity’ and ‘most expensive’ in relation to a solvent
does not necessarily mean ‘no problem’. Secondly, recovery
rates of steroids from water (or plasma) should not be estimated using radioactive tracers, as this will not reveal damage to the steroid. Thirdly, even though ethyl acetate
appears to be a very satisfactory solvent for the elution
of cortisol from solid phase extraction cartridges, it should
not be assumed that it is satisfactory for every other steroid
or compound that one might want to extract.
The most common solvents for elution of steroids from
solid-phase extraction cartridges are methanol or ethanol.
A potential problem with using these solvents is that they
will elute not just free steroids, but also their glucuronides
and sulphates. This may or may not be a problem—
depending on the specificity of the antibody used in the
immunoassay—and should thus be properly validated for
each steroid and species.
7. The amounts of steroid recovered are sometimes too low
In our experience, assay sensitivity has never posed a
problem for measurement of steroids in water. The steep
accurate part of the standard curve in most immunoassays
lies between 10–50 pg per tube. The amounts of steroids
released in pg/g/h are generally well in excess of such values
(Tables 1 and 2). If difficulties are encountered in measuring steroid concentrations, they can be overcome by: leaving the fish for longer in the water (when making static
collections); reducing the water flow rate (in dynamic situations); extracting more water; redissolving the dried-down
extracts in less assay buffer; or changing to a more sensitive
assay procedure. However, by making extracts more concentrated or by increasing assay sensitivity, one runs the
risk of obtaining matrix effects (i.e. introducing compounds
that interfere with the antibodies or labels and yield false
results).
8. The necessity to link the rate of release of the steroid to its
concentration in plasma
Underlying the procedure for measuring steroids in
water is the principle that the concentration of steroid in
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A.P. Scott, T. Ellis / General and Comparative Endocrinology 153 (2007) 392–400
Table 1
Approximate rates of release of stress steroids by different fish species
Steroid
Species
Cortisol
Oncorhynchus mykiss
Oncorhynchus mykiss
Oncorhynchus mykiss
Cyprinus carpio
Rutilus rutilus
Cyprinus carpio
Gasterosteus aculeatus
Archocentrus nigrofasciatus
Perca fluviatilis
Cyprinus carpio
Gobio gobio
Neolamprologus pulcher
Cortisone
Oncorhynchus mykiss
Free steroid release rate (pg/g/h)
Stressor
Reference
Ellis et al. (2004)
Control
Stressed
40
1300
6700
1000
1500
400
2000
690
3500
1800
1700
2100
2560
100–250a
Single handling stress
Repeat handling stress
Confinement for 2 h
Confinement for 2 h
Tagging
Tagging
Stocking and loading density
Handling and confinement
Confinement for 2 h
Ship noise
Ruane and Komen (2003)
Sebire et al., unpublished
Earley et al. (2006)
Wysocki et al. (2006)
Breeding males
Bender et al. (2006)
240
840
Single handling stress
Repeat handling stress
Ellis et al. (2004)
70
170
680
850
1400
850
23
Ellis et al. (2005)
Ellis et al., Fig. 2
Lower et al. (2005)
Values represent ‘ballpark’ estimates of means derived from graphs and tables in relevant publications.
a
Represents release rate of free + glucuronidated + sulphated cortisol.
water equates to the concentration of steroid in plasma, i.e.
there is a direct link between steroid release and the physiology of the fish. It has been pointed out (Scott et al.,
2005) that one of the factors that is likely to affect the rate
of release of steroids into the water is their affinity for sex
steroid binding globulin (SSBG). This has been proposed
as the reason why 17,20b-P is released by female goldfish
at a faster rate than T (Scott and Sorensen, 1994), even
though the concentrations of T in plasma are higher than
those of 17,20b-P (Moriwaki et al., 1991). Data from both
papers is shown replotted in Fig. 1. However, there are
undoubtedly other factors involved, including the lipophilicity of the steroids—that has been shown to influence the
rate of diffusion of drugs across fish gills (Maren et al.,
1968). There is no evidence yet to suggest that the excretion
of steroids via the gills might be an active process in teleost
fish, although specialised cells (Siefkes et al., 2003)—that
actively excrete a sulphated pheromonal sterol into the
water—have been identified in the gills of a primitive vertebrate, the sea lamprey (Petromyzon marinus). Whatever the
mechanism controlling the rate of steroid excretion, as long
as the changes in water follow the pattern of changes in the
plasma (Fig. 1), then there should be no fundamental problem in using water concentrations as a proxy for plasma
concentrations. However, we believe that if measurement
of steroids in water is to be used as a proxy for measurement of steroids in plasma, then it is very important to
establish that such a relationship does exist. This has in
many cases been assumed and/or extrapolated from data
on another species and/or another steroid. In fact, only a
few published papers have so far provided this level of validation of a water steroid procedure (Ellis et al., 2004;
Greenwood et al., 2001; Stacey et al., 1989).
An example of a plasma-water correlation for cortisol
from an unpublished study is shown in Fig. 2. In this study,
two strains of rainbow trout—one selected for high cortisol
response and the other for low cortisol response (Pottinger
and Carrick, 1999)—presented a wide range of cortisol levels to enable a correlation. Eight groups (four from each
strain) of 10 fish were confined in 5 L of water, and cortisol
concentrations in the plasma and water were determined
after 2 h. It must be remembered when examining a plasma-water correlation that a perfect straight-line relationship is unlikely. Plasma steroid concentrations provide a
snapshot at a single time point, whereas water samples represent, to a varying degree, levels over a longer period. Also
plasma concentrations may represent a sample of fish,
whereas water concentrations integrate the release of all
individuals within a population (e.g. Ellis et al., 2004).
9. Metabolism of steroids after release into water
One potential problem with the water steroid procedure
is the metabolism of the steroids whilst in the water. However, to date, the evidence suggests that steroids are reasonably robust, even in static collection mode, and undergo
very little transformation within the short (0.5–2 h) time
frames over which water collection and processing is normally achieved (Ellis et al., 2004; Scott et al., 2005). Water
temperature is likely to be an important factor. In an experiment on the stability of 17,20b-P and Ad in freshwater at
18 °C, both steroids were found to have a half-life of only
6 h (Sorensen et al., 2000). However, in an experiment carried out at 12 °C on naturally-released cortisol in tanks that
had housed fish for a month (and presumably had an active
microbial flora) the average half-life was 16 h (Ellis et al.,
2004). Cortisone, a metabolite of cortisol, is present and
readily measurable in water extracts from rainbow trout
(Ellis et al., 2004). However, it is likely that the conversion
of cortisol to cortisone does not take place in the water, but
while the cortisol is still circulating within the fish (Pottinger and Moran, 1993).
A.P. Scott, T. Ellis / General and Comparative Endocrinology 153 (2007) 392–400
397
Table 2
Approximate rates of release of sex steroids by different fish species
Species
Sex
Steroid
Free steroid release rate (pg/g/h)
Treatment
Reference
Anguilla anguilla
Lythrypnus dalli
Tinca tinca
Rutilus rutilus
Carassius auratus
Gasterosteus aculeatus
Tinca tinca
Carassius auratus
Dentex dentex
Tinca tinca
Rutilus rutilus
Carassius auratus
Carassius auratus
Rutilus rutilus
Rutilus rutilus
Tinca tinca
Tinca tinca
Rutilus rutilus
Carassius auratus
Carassius auratus
Anguilla anguilla
Carassius auratus
Rutilus rutilus
Anguilla anguilla
Anguilla anguilla
Rutilus rutilus
Carassius auratus
Carassius auratus
Oncorhynchus mykiss
Oncorhynchus mykiss
M
M
M
M
M
M
M
M
F
F
F
F
F
M
F
F
M
M
F
M
F
F
F
F
M
M
F
M
Immature
Immature
11-KT
11-KT
11-KT
11-KT
11-KT
11-KT
17,20b-P
17,20b-P
17,20b-P
17,20b-P
17,20b-P
17,20b-P
17,20b-P
17,20b-P
Ad
Ad
Ad
Ad
Ad
Ad
E2
E2
E2
T
T
T
T
T
Melatonina
Melatonin
38
100
250
500
1440
2500
166
200
285
300
1300
2100
3000
9000
120
140
200
600
4000
40,000
6
<5
<5
5
38
125
500
1960
20
15
HCG-injected
Paternal behaviour
GnRHa-injected
Pituitary-injected
HCG-injected
Territorial
GnRHa-injected
HCG-injected
GnRHa-injected
GnRHa-injected
Pituitary-injected
HCG-injected
Natural spawning
Pituitary-injected
Pituitary-injected
GnRHa-injected
GnRHa-injected
Pituitary-injected
HCG-injected
HCG-injected
HCG-injected
HCG-injected
Pituitary-injected
HCG-injected
HCG-injected
Pituitary-injected
HCG-injected
HCG-injected
Nighttime
Nighttime
Huertas et al. (2006)
Rodgers et al. (2006)
Pinillos et al. (2003)
Lower et al. (2004)
Sorensen et al. (2005)
Sebire et al., unpublished
Pinillos et al., 2002
Sorensen et al. (2005)
Greenwood et al. (2001)
Pinillos et al. (2003)
Lower et al. (2004)
Scott and Sorensen (1994)
Van Der Kraak et al. (1989)
Lower et al. (2004)
Lower et al. (2004)
Pinillos et al. (2003)
Pinillos et al. (2003)
Lower et al. (2004)
Scott and Sorensen (1994)
Sorensen et al. (2005)
Huertas et al. (2006)
Sorensen and Scott (1994)
Lower et al. (2004)
Huertas et al. (2006)
Huertas et al. (2006)
Lower et al. (2004)
Scott and Sorensen (1994)
Sorensen et al. (2005)
Ellis et al. (2005)
James et al. (2004)
Five cichlids (including tilapia)
Neolamprologus pulcher
Neolamprologus pulcher
Betta splendens
Five cichlids (including tilapia)
Neolamprologus pulcher
Neolamprologus pulcher
Salaria pavo
M
M
M
M
M
M
M
M
11-KT
11-KT
11-KT
11-KT
T
T
T
17,20b-P
F + S + Gb
60
600
40–100c
700
200
450
100–250c
3600d
Territorial
Territorial
Breeding
Nesting
Territorial
Territorial
Breeding
Breeding
Hirschenhauser et al. (2004)
Oliveira et al. (2003)
Bender et al. (2006)
Dzieweczynski et al. (2006)
Hirschenhauser et al. (2004)
Oliveira et al. (2003)
Bender et al. (2006)
Oliveira et al. (1999)
Values represent ‘ballpark’ estimates of means derived from graphs and tables in relevant publications.
11-KT, 11-ketotestosterone; 17,20b-P, 17,20b-dihydroxypregn-4-en-3-one; T, testosterone; E2, 17b-estradiol; Ad, androstenedione.
a
Melatonin is included because of its potential as a ‘housekeeper’ compound (see text).
b
Represents release rate of free + glucuronidated + sulphated steroid.
c
Data from two sizes of fish (11–4 g).
d
Based on extra data provided by authors—units shown on their graph should have been ‘pg/2 ml’ and average weight of their males was 12.9 g; 76% of
steroid was in sulphated form and 15% in free form.
10. Re-uptake of steroids by fish
The evidence for release of steroids into the water by fish
is strong. The evidence that fish can take up steroids from
the water is equally strong. It was noted in a closed circulation system (Budworth and Senger, 1993), that when T
was injected into one set of fish, it rapidly appeared in plasma of another set of fish. Rainbow trout were shown to
both release and take up 17,20b-P in a way that was amenable to mathematical modelling (Vermeirssen and Scott,
1996). As mentioned above, tench show an ability to take
up steroids from the water that appears to depend strongly
on how avidly the steroids bind to plasma sex steroid
binding globulin (Scott et al., 2005). In fact, fish (see also
a recent study on uptake of E2 and T by the stickleback
(Gasterosteus aculeatus); Maunder et al., in press) appear
to be so efficient at absorbing certain steroids that are added to the water that the final concentrations of these steroids in plasma can reach levels that are 300 times higher
than those in the surrounding water. Paradoxically, this
biomagnification effect calls into question the interpretation of plasma steroid concentrations (not just water steroid concentrations) in experiments where fish are in
close proximity to each other and sharing the same body
of water (especially in recirculation systems). In such
situtations, how much of steroid that can be measured in
398
A.P. Scott, T. Ellis / General and Comparative Endocrinology 153 (2007) 392–400
should be reported as ‘apparent’ or ‘uncorrected’ release
rates.
11. Using fish of different sizes
Fig. 1. Concomitant changes in rates of release and plasma concentrations of T (n) and 17,20b-P (s) in HCG-injected female goldfish to
illustrate that, although the ratios between the two steroids are totally
different in plasma and water (presumed to be due to the much higher
affinity of T for plasma sex hormone binding globulin), the pattern of
release of both steroids is nevertheless the same as the pattern of change of
plasma concentrations.
Fig. 2. Mean plasma cortisol concentration at end of a 2 h confinement
period (n = 10 fish per point) plotted against cortisol release rates during
the 2 h confinement period for rainbow trout. HR, selectively bred for a
high cortisol response; LR, selectively bred for a low cortisol response.
rs = 0.91, p = 0.02.
On purely theoretical grounds, the smaller the fish, the
higher the rate of steroid release due to a higher ratio of gill
surface area to body mass. This has not yet been the subject
of focussed investigation—but was found to be a confounding factor when using total steroid release rates to
delineate social status and behaviour in males (‘breeders’
and ‘helpers’) that had large differences in body mass
(Bender et al., 2006). When calculated as pg/fish/h, there
were no differences in excretion rate of total T, 11-KT
and cortisol between the two types of males. However,
when calculated as pg/g/h, the rate of excretion by the
smaller fish was higher in inverse ratio to the difference in
mass between the two groups. These results highlight a
major difficulty in interpreting steroid release rates and
underline the necessity of finding a ‘normaliser’ for steroid
concentrations in water.
12. Suggested validation steps for water steroid procedure
As a conclusion to this review, we provide a checklist for
validating a procedure for measuring a steroid in water. In
our opinion, the temptation to extrapolate results from one
species to another (or from one steroid to another) should
be resisted. The main questions to be answered are:
Is one measuring the most appropriate form of steroid
(e.g. cortisol vs cortisone; T vs 11-KT)?
Is the steroid stable during short-term storage (frozen
storage if necessary) and extraction?
Is the recovery rate stable?
Is there evidence for undue non-specific interference in
the assay (i.e. false positives from fish-free water extracts)?
Is the immunoassay specific (i.e. are there perhaps other
steroids in the water that might cross-react with the
antibody)?
Are steroid concentrations related to those in plasma
(linear or curvilinear)?
Are the results biologically meaningful?
Acknowledgments
plasma is made by the fish itself and how much by the other
fish in the tank? In the many cases where 11-KT is found in
the plasma of females or E2 in the plasma of males, in how
many of these cases might this be due to ‘fish-to-fish’ transfer of steroids rather than by internal biosynthesis?
To date, no attempts have been made, in any published
studies, to take into account the effects of steroid re-uptake
and metabolism on recorded values for steroid release rate.
Also, in only a very few studies have values been corrected
for losses incurred during extraction. Thus most values
shown in Tables 1 and 2 are almost certainly underestimates of the true release rate. It could be argued that they
We acknowledge financial support from Defra Animal
Welfare and Health Division and the European Union
(6th Framework Project No 501984: WEALTH). We thank
Professor Adelino Canario, Faro University for helpful
critical comments on the manuscript.
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