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 396 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). 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