gy, Part C 143 (2006) 275–283www.elsevier.com/locate/cbpc

Comparative Biochemistry and Physiolo

Pharmacokinetics of morphine in fish: Winter flounder (Pseudopleuronectesamericanus) and seawater-acclimated rainbow trout (Oncorhynchus mykiss)

Nathalie C. Newby a, Paula C. Mendonça b, Kurt Gamperl b, E. Don Stevens a,⁎

a Department of Integrative Biology, University of Guelph, Ontario, Canada N1G 2W1b Ocean Sciences Centre, Memorial University of Newfoundland, St. John's, NF, Canada A1C 5S7

Received 12 October 2005; received in revised form 3 February 2006; accepted 6 March 2006Available online 2 May 2006

Abstract

We made a single intraperitoneal (IP) injection of morphine sulfate (40mg/kg) into winter flounder and seawater acclimated rainbow trout at10°C and then followed its disposition by measuring the change in plasma morphine concentration for 100h using a morphine specific ELISA.Disposition also was followed for 6h after a single IV injection of 7.5mg morphine sulfate in winter flounder. Plasma morphine reached amaximum within an hour post-injection IP and then decreased in a bi-exponential fashion with a rapid distribution phase followed by a slowerelimination phase. The disposition was slower in flounder than in trout even though the fish were held at the same temperature. For example,plasma clearance was 76mL h− 1 kg− 1 in the flounder but was almost twice as much in the trout (153mL h− 1 kg− 1) and mean residence time was27.9h in the flounder but was 7.0h in the trout. The present study is the first comprehensive pharmacokinetic analysis for any analgesic in anectotherm, and our results show that: 1) significant intra-specific variation exists between fishes: and 2) the disposition of morphine in fish isapproximately one order of magnitude slower than it is in mammals. These differences may be due in part to mass specific differences in cardiacoutput.© 2006 Elsevier Inc. All rights reserved.

Keywords: Analgesia; Analgesic; AUC; AUMC; Clearance; Fish; Kinetics; Morphine; Pain; Teleosts

1. Introduction

There is a considerable body of literature, especially in thenon-peered reviewed or gray literature, concerning pain in fishand what should be done about it. However, there is very littleexperimental data concerning the efficacy or the pharmacoki-netics of analgesics in any fish species. Moreover, it is wellknown that extrapolation of pharmacokinetic or pharmacody-namic parameters across species is not appropriate (Riviere,1999). The present study is one of a series our laboratory isundertaking to provide data that may be useful to organizationswriting guidelines regarding the use of fish in research andteaching. Here we provide the first pharmacokinetics data for ananalgesic in fish. We chose morphine because it is the standardagainst which all other analgesics are compared.

⁎ Corresponding author. Tel.: +1 519 824 4120x52137; fax: +1 519 767 1656.E-mail address: dstevens@uoguelph.ca (E.D. Stevens).

1532-0456/$ - see front matter © 2006 Elsevier Inc. All rights reserved.doi:10.1016/j.cbpc.2006.03.003

The only other study that has any data concerning thepharmacokinetics of analgesics in fish is that of Jansen andGreene (1970). They transferred goldfish to water containing10mg/L morphine and reported that “absorption was complet-ed in approximately 15min, no further uptake occurring duringthe next three hours”. That is, the concentration in the fish'stissues (9.7mg morphine/kg fish) equilibrated so that it wasessentially the same as in the water (10mg morphine/L water).Moreover they reported that when the fish (now containingmorphine) was placed into a fresh container, the efflux ofmorphine from the fish occurred at the same rapid rate. This isan extraordinary result given that morphine is a relatively largemolecule (molar mass 285). Moreover, it is a charged moleculeat physiological pH values and exists mostly (64%) as a cationat physiological pH (7.85 in flounder at 10 °C); with a positivecharge on the tertiary amine (HOBNH+) having a pKa1 valueof 8.10 at 10 °C (Stevens, unpublished). The pKa2 for the lossof a proton from the phenolic hydroxyl group at carbon 3 is9.84 at 10 °C (Stevens, unpublished). Further, we have

276 N.C. Newby et al. / Comparative Biochemistry and Physiology, Part C 143 (2006) 275–283

replicated the experiment of Jansen and Greene (1970), butcannot reproduce their results. Instead, based on blood andwater morphine measurements, we show that uptake across thegills takes days, not minutes (Newby, Wilkie, and Stevens,unpublished). Jansen and Greene did not measure morphine inthe blood. Insofar as we know, no other study has measuredmorphine concentrations in fish blood.

Although the metabolism of morphine differs somewhat indifferent mammals, the two major metabolites are morphine-3-glucuronide (M-3-G) and morphine-6-glucuronide (M-6-G). M-6-G has analgesic effects and can account for a significantportion of morphine's analgesic action in mammals (Milne etal., 1996). M-3-G does not have antinociceptive effects, butresults in hyperalgesia in some mammals (Milne et al., 1996).The metabolites are excreted by the kidney. Insofar as we know,the metabolites of morphine have never been measured in fish,but we have no reason to believe they would differ in anysubstantive way.

In the present study, we examined the distribution andelimination of morphine in winter flounder (Pseudopleuronectesamericanus) and seawater-acclimated rainbow trout (Oncorhynchusmykiss).Weadministeredasingledoseofmorphinesulfate into theperitoneal cavity, took repeated blood samples at different timeintervals, and measured the concentration of plasma morphineusing a morphine specific direct enzyme-linked immunosorbantassay. Intuitively,weexpectedthat thechange inplasmamorphineafter a single intraperitoneal (IP) injection might involve threephases with different time courses: i) uptake from the IP site toplasma, ii) exchange between plasma and all other tissues, and iii)elimination from the animal.However, absorptionwas extremelyrapidbecauseour IPcannulawas locatedbetween the liver and thebody wall. A characteristic property of drug elimination after asingle injection in mammals is that there are two majorcomponents to the clearance kinetics: a fast phase and a slowphase (Riviere, 1999)—this model turned out to best fit our datafor fish as well.

We chose the IP route to administer morphine for pragmaticreasons. The purpose of our study has important implicationsfor guidelines for fish care. If morphine ever is to be used in fishduring or after a procedure, then the most reasonable route ofadministration is IP. Cannulation with IV administration isimpractical in small fish and extremely difficult in many fishspecies. The commonest surgical procedure where morphinemight be implicated is in the surgical implantation of telemetrydevices. It would be inappropriate to insist that researchersimpose further surgical trauma just to administer an analgesic.However, we also used the IV route in winter flounder in orderto estimate some pharmacokinetic parameters.

2. Materials and methods

2.1. Animals

Ethical approval was obtained from the Animal CareCommittees both at the University of Guelph and at MemorialUniversity of Newfoundland. Winter flounder (P. americanus)were caught by divers using a hand net in Conception Bay

(Newfoundland) at a depth of 4–6m and transported to theOcean Sciences Centre (OSC, Memorial University of New-foundland). These fish initially were held in a 2×2×0.5m deeptank supplied with 4 °C seawater (pH 8.0 to 8.1) at 10L/min for3weeks. Thereafter, the temperature was raised to 10 °C over aperiod of 2–3weeks. One week prior to experimentation, thefish were transferred to a holding tank (2×2×0.3m) in theexperimental room where they were held at 10 °C for anadditional week. The flounder were fed diced herring twice aweek (ca., 1% body weight) but were fasted for 18h prior tosurgery.

Seawater-acclimated rainbow trout (O. mykiss) wereobtained from a commercial aquaculture operation (Long IslandResources, Bay D'Espoir, NL, Canada) and held for approx-imately 9months at the OSC in flow through tanks (3mdiameter×5m deep) receiving seawater at 10 °C. They were fedcommercial trout pellets 3 times per week but fasted 18h priorto surgery.

2.2. Surgery

Fish were anaesthetized in seawater containing MS-222(0.25g/L). Once anaesthetized, fish were weighed on a panbalance (Ohaus Scout accurate to 1g) after removing as muchsuperficial water as possible, and then transferred to a surgicaltable, where their gills were irrigated with chilled (∼4 °C) andoxygenated seawater containing 0.1g/L MS-222.

2.2.1. Flounder cannulationsWe made a 0.5cm long incision, just below the lateral line

about one-third of the animal's length from the tail and retractedthe skin and underlying muscle tissue to expose the post-cardinal vein which lies between the hemal arches. We inserteda heparinized cannula (PE 50, Clay Adams, 80cm long, volume0.2mL) with indwelling 14-gauge piano wire into the vessel.Finally, after removing the indwelling wire, and pushing thecannula approximately 8cm anteriorly into the vein, the incisionwas sutured, and the cannula was filled with heparinized saline(0.9% NaCl with 100IU mL− 1 heparin) and sutured to the fish'sdorsal surface at two locations. There was minimal bleedingduring cannula placement, and the cannula was kept open byperiodically flushing with saline until the experiments began.

We used an indwelling cannula in the peritoneal cavity toinject the morphine because these fish are very sensitive tohandling and we wanted to minimize any handling effectsduring the morphine or control injections. To implant the IPcannula, the animal was placed blind-side up, a hole in the bodywall was made at the distal edge of the peritoneal cavity usingan 18 gauge needle, and the IP cannula (PE 50, Clay Adams,80cm long, volume 0.2mL) was inserted into the anteriorportion of the peritoneal cavity so that the cannula tip wasbetween the liver and cavity wall. Finally, the cannula wassutured at a position close to where it entered the peritonealcavity, passed to the eyed-side (dorsal side) of the animalthrough a hole in the body musculature made near the ventral finwith a 14 gauge needle, and sutured twice more to the skin onthe eyed side.

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Upon completion of surgery, the flounder was placed in theexperimental chamber (a 50×30×25cm opaque plastic con-tainer supplied with aerated 10 °C seawater, at 5L min− 1 andfilled with sand to a depth of 5cm), and recovered by flushingthe gills with seawater for about 2–5min.

2.2.2. Trout cannulationsThe trout were fitted quickly (within approximately 1 min)

with a dorsal aortic cannula (PE 50, 80cm long) using themethod of Smith and Bell (1964), then placed on their left sideso that the IP cannula could be inserted. To insert the IP cannulainto the trout, a hole was made in the body wall immediatelybehind the pectoral fin, the cannula (PE 50, 80cm long) waspushed approx. 8cm posteriorly into the cavity so that thecannula tip was between the liver and cavity wall, and thecannula was sutured to the skin close to where it entered theperitoneal cavity and again near the fish's dorsal fin. Aftersurgery, trout were placed in individual ‘black boxes’ (70cmlong×12cm wide×15cm deep) supplied with aerated 10 °Cseawater, and recovered in a similar fashion as flounder.

2.3. Experimental procedures

Both the flounder (n=7; 564.6±81.6g) and trout (n=7;1002.7±68.8g) were allowed to recover for at least 18 h prior toexperimentation. An initial blood sample (0.25mL) was takenfrom each fish so that pre-injection levels of morphine could beassessed. Thereafter, morphine sulfate (15mg/mL) was injectedinto the IP cannula of each fish at a nominal dose of 40mg/kgbody weight, and the IP cannula was flushed with 0.2mL ofsaline to ensure that all the morphine was delivered to the fish.Morphine was injected with a 3mL syringe (average dosevolume 1.5mL). The potential error in dosing was estimated byrepeated weighing of nominal 1.5mL samples delivered with a3mL syringe: mean volume 1.50mL, S.D.=0.0099, n=20,range from 1.48 to 1.52. The potential error in dosing was about2%. Blood samples (0.25mL) were then obtained from all fishat 0.25, 1, 4, 10, 40, and 100h post-injection, immediatelyplaced into 1.5mL centrifuge tubes, containing 25μL heparin,and placed on ice. Blood samples were subsequently centri-fuged for 5min at 6000rpm (Eppendorf 5415 C), the plasmawas transferred to cryovials, and stored at −80°C until analysis.At the end of the experiment, all fish were euthanized byimmersion in MS-222 (0.3g/L) and dissected to confirm theposition of the cannulas and to check for bleeding at cannulationsites. In a separate experiment on 4 winter flounder, dispositionwas followed after a single IV injection of 7.5mg morphinesulfate. The intention here was to get a better estimate of theparameters for distribution, so in this case blood samples weretaken at 1, 1.5, 2, 3, 4, 5, and 6h post-injection.

2.4. Sample analysis

We analyzed blood samples using a morphine specific directenzyme-linked immunosorbant assay (ELISA) kit (BQ 213-0192; Bio-Quant Inc., San Diego, USA). We made a workingstock solution of 1000ng/mL morphine sulfate in water from a

stock of 0.5mg/L morphine sulfate in methanol then produceda standard curve in plasma using morphine sulfate concentra-tions of 0, 2.5, 5.0, 10.0, 25.0, 50.0, 100.0ng/mL. Plasmasamples were diluted so that concentrations were near theoptimum for the assay. We plated our plasma samples andplasma standards following the ELISA Kit procedures providedby BioQuant. Standard curves for the ELISA were fitted to alogistic dose response curve {Y=a+b / (1+ (x /c)d)} using non-linear least-squares regression with TableCurve 2D v3 (JandelScientific Software). Adjusted r-squared values for these fitswere >0.99.

2.5. Chemicals

Morphine sulfate (15mg/mL, injection USP, for intramus-cular, subcutaneous, or intravenous administration) was pur-chased from Sabex® (Boucherville, QC, Canada) and MS-222(TMS or tricaine methane sulfonate) was purchased fromSyndel Laboratories (Vancouver, B.C., Canada).

2.6. Data analysis

As with most pharmacokinetic analyses, we assumed that theprocesses that governed disposition of plasma morphine werefirst order (Ritschel, 1992). Plasma morphine concentrationsversus time data were analyzed using non-linear least-squaresregression analysis using GraphPad Prism 4.03 (GraphPadSoftware Inc.). The fitting program was initialized withparameter values estimated by curve-stripping. A two-compart-ment open model with 1/y2 weighting proved to be the best fitfor our data:

Plasma ½morphine sulfate� ¼ A⁎exp−ða⁎timeÞþ B⁎exp−ðb⁎timeÞ

and the adjusted r-squared values for all fits were >0.9; mean r-squared value was 0.98. Fits to the flounder IV data wereconstrained using the mean β from the flounder IP study.

There are many different equations used to calculatepharmacokinetic parameters. Thus we briefly describe theforms we used. First, the disposition curves (plasma morphineconcentration versus time) of the two species were comparedusing a global nonlinear bi-exponential model with two speciesand 7 replicates for each species. Then the curve was fit to eachfish separately to calculate individual equation parametervalues. The individual concentration versus time values andthe individual β values were then used to calculate AUC andAUMC.

The area under the concentration versus time curve from zeroto infinity (AUC∞) and its first moment (area under theconcentration× time versus time curve from zero to infinityAUMC∞) were calculated using two methods for each fish.First, from the observed data points (AUC∞area, AUM-C∞area), and second from the parameters of the exponentialequation (AUC∞ expo, AUMC∞ expo). AUC∞ area andAUMC∞area to the final concentration time point (Cn) werecalculated by the linear trapezoidal rule using the observed data

Fig. 1. Plasma morphine sulfate disposition in winter flounder after a single IPinjection of 40mg/kg morphine sulfate at time zero. (A) Linear scale with best fitregression line showing raw data for 7 flounder with different symbols fordifferent fish. (B) Logarithmic scale with the best fit regression line; points aremeasured plasma levels (mean±S.E.M.). Scales are the same as those used fortrout (Fig. 2) to facilitate comparisons.

278 N.C. Newby et al. / Comparative Biochemistry and Physiology, Part C 143 (2006) 275–283

points and then extrapolated to ∞ by adding the value from Cn

to ∞. The concentration at time zero (C0) was zero for the IPcalculations.

AUC obs area ¼X

ððtiþ1−tiÞ=2Þ⁎ðCi þ Ciþ1Þ fromt ¼ 0 to t ¼ n−1

AUClarea ¼ AUC obs areaþ Cn=b

and

AUMC obs area ¼X

ððtiþ1−tiÞ=2Þ⁎ððCi⁎tiÞ þ ðCiþ1⁎tiþ1ÞÞfrom t ¼ 0 to t ¼ n−1

AUMCl area ¼ AUMC obs areaþ ðCn⁎tn=bÞ þ ðCn=b12ÞWe also report these areas calculated using the parameters

from the exponential equation:

AUCl expo ¼ A=aþ B=b

and

AUMCl expo ¼ A=a12þ B=b12

The half-life for distribution was calculated as t1/2dist= ln(2)/α and for elimination as t1/2elim=ln(2)/β. Mean residence time(MRT) was calculated from the observed data points for eachfish using MRT∞area=AUMC∞area/AUC∞area. Clearancebased on the observed values was calculated as Cl∞area=D/AUC∞area and is presented as a mass specific value.

The pharmacokinetic values for the two species werecompared using one-way ANOVAs. The criterion for statisticalsignificance was p<0.05.

The IV injection data were used to estimate the volumes ofdistribution by calculating the microconstants as given inRiviere (1999). Vdss is the volume of distribution at steady state,that is, the volume of distribution at the time when the rate ofelimination equaled the rate of distribution. Total halftime,t1/2total, reflects the halftime for both distribution andelimination processes.

C0=A+Bk21= (A⁎β+B⁎α)/C0

k10=α⁎β/k21k12=α+β−k21−k10Vc or V1=central compartment volume=D/C0

Vdss=Vc⁎ (k12+k21)/k21V2=peripheral compartment volume=Vdss−Vc

Cl=k10⁎Vc

t1/2total= ln(2)⁎Vdss/(k10⁎Vc)

3. Results

3.1. IP study

Morphine was not detected in the plasma prior to injection,but increased rapidly after the IP injection, reaching maximumvalues in all fish within an hour. Maximum plasma values

varied considerably between fish, ranging from 31 to 87mg/L inflounder and from 46 to 87 in trout, even though all fish wereinjected with the same dose (nominally 40mg/kg fish).Maximum plasma [morphine] was not correlated with fishweight (r2 =0.00).

Plasma [morphine] decreased rapidly during the first dayafter the single IP injection in both species (Figs. 1 and 2). Atwo-component model best described the decrease andparameter values for the best fit are given in Table 1. As canbe seen from Figs. 1 and 2 (where the scales on the ordinate arethe same for both species to facilitate comparison), morphinewas eliminated much more rapidly in the trout than in theflounder. Indeed, the plasma concentrations in the trout werestatistically significantly less than in the flounder at 10, 40, and100h post-injection. This point is illustrated in Fig. 3 where themeans for both species are plotted showing that there is muchoverlap between species during the distribution phase but thereis almost no overlap in the elimination phase.

A nonlinear model using data from both species only duringthe distribution phase (0.25 to 10h), and using one exponentialcomponent, showed that the species difference during thisperiod was not statistically significant (F2,46=2.097, p=0.13).In contrast, the same nonlinear model using data from bothspecies during the elimination phase (10–100h) showed that thespecies difference was highly significant (F2,38 = 34.26,p<0.0001).

Time (hours)

0 20 40 60 80 100

Plas

ma

mor

phin

e (m

g/L

)

10 -1

10 0

10 1

Plas

ma

mor

phin

e (m

g/L

)

0

20

40

60

80A

B

trout

Fig. 2. Plasma morphine sulfate disposition in seawater acclimated rainbow troutafter a single IP injection of 40mg/kg morphine sulfate at time zero. (A) Linearscale with best fit regression line showing raw data for 7 trout with differentsymbols for different fish. (B) Logarithmic scale with the best fit regression line;points are measured plasma levels (mean±S.E.M.). Scales are the same as thoseused for flounder (Fig. 1) to facilitate comparisons.

Time (h)

0 20 40 60 80 100

Plas

ma

mor

phin

e (m

g/L

)

0.01

0.1

1

10

100

flounder

trout

rabbit

Fig. 3. Plasma morphine sulfate disposition in winter flounder relative toseawater acclimated rainbow trout after a single IP injection of 40mg/kgmorphine sulfate at time zero. Species differences were minimal in the rapiddistribution phase where the data for flounder and trout overlapped considerablyafter the single IP injection. Morphine was eliminated more slowly in theflounder (circles) relative to the trout (triangles) where there is minimal overlapbetween data points for the two species. Dashed line shows disposition ofmorphine in the rabbit after a single IV injection (data from el-Sayed and Hasan,1990).

279N.C. Newby et al. / Comparative Biochemistry and Physiology, Part C 143 (2006) 275–283

The distribution phase (α) was very rapid, whereas theelimination phase (β) was slow. For example, the half-time fordistribution was less than 3h for both species, whereas the half-time for the elimination phase (t1/2elim=ln(2)/β) was about 34hfor the flounder and 14h for the trout (p=0.026) (Table 1).

Table 1Pharmacokinetic parameters for the disposition of plasma [morphine sulfate]after a single IP injection of 40mg/kg in winter flounder and in SW acclimatedrainbow trout

Units Flounder n=7 Trout n=7 p

Mass g 565±76 1003±64 0.001A mg/L 63.5±8.0 82.5±8.5 nsα per hour 0.434±0.083 0.500±0.032 nsB mg/L 8.76±2.19 4.08±1.23 nsβ per hour 0.0274±0.0054 0.0527±0.0051 0.008D=dose mg 22.1±3.0 39.4±2.4 0.001D/kg mg/kg 39.0±0.3 39.3±0.1 nst1/2elim h 34.1±7.3 13.9±1.1 0.026t1/2dist h 2.20±0.46 1.43±0.09 nsAUC∞expo mg h/L 513±72 245±36 0.009AUC∞area mg h/L 583±74 290±37 0.007AUMC∞expo mg h2/L 18,600±5970 1800±378 0.023AUMC∞area mg h2/L 18,600±5990 2055±345 0.026MRT∞area h 27.9±6.6 7.0±0.6 0.012Cl∞area mL h−1 kg−1 75.6±10.3 153±20 0.008

The last column is the probability of a species difference, where p<0.05 wasconsidered statistically significant.

Fig. 4. Plasma morphine sulfate disposition in winter flounder after a single IVinjection of 7.5mg morphine sulfate at time zero. (A) Linear scale with best fitregression line showing raw data for 4 winter flounder with different symbols fordifferent fish. Inset shows the same figure with an expanded time scale andincludes data at 40h from the IP study. (B) Logarithmic scale with best fitregression line, points are measured plasma levels (means±S.E.M.). Inset showsthe same figure with an expanded time scale and includes data at 40h from the IPstudy.

Table 2Pharmacokinetic parameters for the disposition of plasma [morphine sulfate]after a single IV injection of 7.5mg in winter flounder (Pseudopleuronectesamericanus)

Units Flounder n=4

Mass g 433±16A mg/L 104±16.1α per hour 1.27±0.09B mg/L 6.95±0.53β per hour 0.0274±0.005C0 mg/L 111±15.7D=dose mg 7.50±0.0D/kg mg/kg 17.4±0.65AUC∞area mg h/L 340±12AUMC∞area mg h2/L 11,850±42MRT∞area h 29.3±1.0k21 per hour 0.117±0.020k10 per hour 0.327±0.043k12 per hour 0.855±0.104V1=Vc L 0.170±0.024Vdss L 1.42±0.137V2 L 1.25±0.115t1/2total h 19.1±1.23

280 N.C. Newby et al. / Comparative Biochemistry and Physiology, Part C 143 (2006) 275–283

Mean residence time after the IP injection was about fourtimes longer in the winter flounder than in the trout (Table 1)because of the slower elimination phase in the flounder. Wedid not have adequate data to calculate the absorption phaseusing a 3 phase model. However, preliminary calculationsusing a 3 phase model suggested that the absorption phase hada coefficient of 16.1—a value 37 times greater than thedistribution coefficient and more than 600 times greater thanthe elimination coefficient.

The AUC and AUMC calculated from the terms in the fittedequation (∞ expo) were similar to those calculated from theobserved values (∞ area) and these areas are substantially lessfor the trout than the flounder largely because the plasmamorphine values at 4, 10 and 100h were lower in the trout.Finally, the clearance of morphine from the plasma was almosttwice as fast in the trout (153mL h− 1 kg− 1) as that in theflounder (76mL h− 1 kg− 1).

3.2. IV study in flounder

Plasma morphine decreased rapidly after the single IVinjection (Fig. 4), and the parameter values and calculatedmicroconstants are given in Table 2. The AUC and AUMCvalues are less than those in the IP study because the dose wasless (17.4mg/kg and 39.0mg/kg respectively). However,MRT∞area was 29.3h, a value very similar to that measuredin the IP study (27.9h). The Vdss was 1.42L; the V2 componentwas about 7 times larger than Vc, the central compartment.

4. Discussion

Our study is the first comprehensive pharmacokinetic studyfor any analgesic in any ectotherm and shows first that there aresubstantial differences between fish species. This is important towriters of guidelines for the use of fish in research and teaching

because it means that species differences must be taken intoaccount. Our results also show that the disposition of morphineis about one order of magnitude slower in trout than inmammals (Fig. 3).

In both fish species, the pharmacokinetic data showedconsiderable variation between fish subjects. This variation wasnot related to fish size, and although we do not know the age ofthe fish or record their gender, none of the fish wasreproductively mature. In humans, there are pharmacodynamicgender differences for morphine (i.e., the analgesic effect at thesame plasma morphine concentration is different) but nodifferences in pharmacokinetics (Sarton et al., 2000). On theother hand, age does influence morphine pharmacokinetics inhumans; elderly humans show decreased morphine clearancewith a trend to a smaller volume of distribution. Largeunexplained variation between individuals is characteristic ofstudies on analgesics, especially in pharmacodynamics (e.g.,Parton et al., 2000) and this would be an interesting avenue topursue in fish.

Our AUC and AUMC values cannot be directly comparedwith the mammalian studies because their calculation is afunction of the injected dose. However, the fact that both theseterms were less in the trout than in the flounder indicates thatelimination was faster in the trout and confirms that substantialintra-specific differences exist between fishes. This result hasconsequences for writers of guidelines for the use of analgesicsin fish—pharmacokinetic data will have to be collected on anumber of fish taxa before valid guidelines can be written. Allof our experiments were done in the same room using the samewater supply at the same temperature—thus the differencesreflect actual species differences. We do not know the reason forthese differences, but speculate that they may be related in partto the higher metabolic rate and/or cardiac output (Fig. 4A) oftrout relative to flounder. Disposition will be a function of bloodsupply to the tissues involved in metabolism (liver) andexcretion (kidney and/or liver). It also will depend on thecapacity of the liver to convert the morphine into products thatcan be excreted—probably morphine glucuronides, and thecapacity of the kidney and/or liver to excrete these metabolites.In mammals, most of the morphine is excreted by the kidneys asglucuronides, but the relative proportions of metabolites differin different mammals. For example, in rats 19% is excreted asM3G, whereas in humans 55% is excreted as M3G. Insofar aswe know, the metabolites have not been measured in fish.

The blood sample sites were different in the two species:postcardinal vein in flounder and dorsal aorta in trout. It isunlikely that this difference in sample site accounts for themarked difference in elimination rates that we observed in thetwo species. First, the disposition is described by the slope ofthe line (i.e., β in Tables 1 and 2). If our sample site in theflounder was upstream of the organ that eliminated themorphine, although all of the values would be slightly higher,the slope would be the same. Similarly, if the sample site wasdownstream of the organ that eliminated the morphine, thenall of the values would be slightly lower, but the slope wouldbe the same. Second, it is extremely unlikely that movementof morphine across the gills into the water could account for

Cle

aran

ce (

mL

/min

*kg)

1

10

100

f

t

rrr

rr

c

rbrbrb

rbrb ddd

ddss

hh

hhhhhhh

Cardiac output (mL/min*kg)

10 15 20 30 40 60 80 100 150 200

MR

T (

h)1

2

4

68

10

1520

30 f

t

r

c

rbdd

d

dd

hh

A

B

Fig. 5. Pharmacokinetic parameters in relation to cardiac output for fish (presentstudy) and various mammals. Letters inside the symbols indicate the species:f= flounder, t= rainbow trout, h=human, s=sheep, c=cat, rb=rabbit, d=dog,r= rat. The solid line in A is the unity line, the dashed line is the least squaresregression line y=0.0233⁎x^1.57. Scales for cardiac output, clearance, andMRT are logarithmic. Routes of administration for flounder and trout are IP andfor mammals are IM or IV. Values of cardiac output are from: flounder (Joaquimet al., 2004, Mendonça, P. and Gamperl A.K. unpublished); trout (Gamperl et al.,1994; Thorarensen et al., 1996; Brodeur et al., 2001); man, rabbit, dog, and rat(Popovic et al., 2005); cat (Groom and Rowlands, 1958) sheep (Stowe andGood, 1961). Values of pharmacokinetic parameters are from: flounder and trout(present study); cat, dog, and human (from Table 2 in Taylor et al., 2001) rat,rabbit, dog, sheep, and human (from Table 9 in Milne et al., 1996); rabbit (el-Sayed and Hasan, 1990); rat (Letrent et al., 1998).

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the difference. Morphine movement across the gills ingoldfish is less than 0.003% in 4h (Newby, Wilkie, andStevens, unpublished).

The Phish-Pharm database has 2342 entries of which 23 arefor flounder (not winter flounder) (Reimschuessel et al., 2005).Of the flounder entries, there are three studies with pharmaco-kinetics data, two are for pollutants (PCBs and BAP) and onefor oxytetracycline. For the oxytetracycline study the authorsstate “the scarcity of time points prevented us from conductingkinetic analysis” and in their table of parameter values they didnot give any data for flounder (“parameters were not determineddue to insufficient data points”) (Chen et al., 2004). However,the authors of the database estimated the t1/2 from the raw datafor summer flounder as 155h. There are 13 entries foroxytetracycline disposition in rainbow trout; mean t1/2 was151±32h (mean±S.E.M.), essentially the same as that for thesummer flounder. Moreover, the data from the 13 studies fortrout showed that the t1/2 for oxytetracycline did not scale withbody mass (r2 =0.06) or with water temperature (r2 =0.19). Thestudies for PCBs and BAP do not give any parameter values thatpermit comparisons between flounder and trout because theydid not sample blood. Thus, the Phish-Pharm database,although extensive, does not provide examples that mightgive some insight into mechanistic explanations for thedifference between winter flounder and rainbow trout withrespect to our observation on the disposition of morphine.

The disposition of morphine was considerably slower in fishthan reported for mammals (Fig. 5). The plots in Fig. 5 compareour results for flounder and trout with those of mammals. Ineach case, the pharmacokinetic variable is plotted versus thecardiac output. Cardiac output is used on the abscissa becauseclearance is a measure of the amount of blood cleared per unittime and morphine disposition could be perfusion limited, atleast in part.

Clearance scales, at least in part, with cardiac output; for ourtwo species of fish and across all species in general (r2 forclearance=0.94) (Fig. 5A). However, there are importantspecies differences that are not accounted for by differencesin cardiac output. For example, the cat and rabbit are similar insize and have similar cardiac output values and metabolic rates,but the clearance rate is about five times higher in the rabbit. Inthe flounder, trout, and cat clearance ranges from 9% to 18% ofcardiac output, whereas in the rabbit and rat it ranges from 40%to 70% of cardiac output. This is illustrated in the figure becausethe points for the rabbit and rat are much closer to the unity linethan those for the flounder, trout, and cat.

MRT scales inversely with cardiac output, both for our twospecies of fish and across all species in general (r2 forMRT=0.93) (Fig. 5B). However, cardiac output for thesespecies comparisons is strongly correlated with metabolic rate.Moreover, for all of the points in Fig. 5, the cardiac output andthe pharmacokinetic values are taken from different studies. Itwould be fruitful to measure both the disposition of morphineand cardiac output simultaneously in a variety of species toexamine this relation more closely. The fact that the relationseems to fit across all species suggests that the difference indisposition between fish and mammals cannot be attributed to

differences in phylum or to a difference in respiratory medium.Rather, it seems to suggest that the disposition is a function ofcardiac output and/or metabolic rate. However, there are otherfactors that determine disposition that are important. The rate ofhepatic biotransformation and enterohepatic recirculation surelydiffers between these species. Enterohepatic recirculation orrecycling of morphine is important in mammals. A smallfraction of the biotransformed morphine is excreted in the bile,biotransformed by bacteria in the gut back into morphine,absorbed by the gut and hence recycled. This possibility has notbeen studied with regards to morphine in fish, and in fact is verypoorly studied in fish for any substance. It is unlikely thatpercentage body fat could account for the huge differencesbetween mammals and fish, but could play a role in explainingdifferences between fish species.

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Similar arguments regarding scaling of clearance and MRThave been made by Riviere (1999), although he plotted thevariables against body weight. Plotting against body weight isappropriate when making comparisons within mammals, but isnot appropriate when comparing mammals and ectotherms,such as fish, that have much lower mass specific metabolic ratesand cardiac outputs than mammals.

It must be pointed out that there are some limitations in ourstudy. In particular, the parameter values calculated fromAUC∞area and AUMC∞area are more meaningful than valuescalculated from the exponents of the fitted curve. There areequations available that are used to calculate clearance andmean residence time using the equation exponents, but thesewill be less accurate than the values we report here calculatedusing AUC∞area and AUMC∞area. The half-times reportedin Table 1 must be viewed with some caution because they arecalculated using the exponents of the bi-exponential fittedcurves. However, no matter how we calculate the t1/2elim (usingmean concentration values, calculating from mean β values orusing median values) the t1/2elim for morphine was much longerthan for any mammal.

Insofar as we know, there are no other comprehensive studiesof pharmacokinetics of morphine in any ectotherm. However, ithas been reported that “the onset and offset of the analgesiceffects of morphine appear to be much slower in amphibiansthan in rodents” (Stevens and Kirkendall, 1993). In mammals,morphine is an effective analgesic at plasma concentrations ofabout 1μM (about 0.285mg/L). Admittedly, the range of steadystate concentrations associated with analgesia in cancer patientsis broad (0.016 to 0.346mg/L) (Gustein and Akil, 2001).Moreover, the antinociceptive effect of morphine lags behindthe plasma concentration. In rats the plasma concentration fallsto about half-maximum in 60min, whereas the antinociceptiveeffect takes about 120min to fall to half-maximum (Letrent etal., 1998). In our experiments, the plasma concentration wouldbe above the antinociceptive effective level in mammals forabout 53h after a single injection (estimated by interpolationbetween the points at 40 and 100h). In the frequently citedpapers on pain in fish by Sneddon and colleagues, a dose of300mg/kg was used (Sneddon, 2003: Sneddon et al., 2003), 7.5times the dose used in our study. If the analgesic efficacy ofmorphine is similar in trout as it is in mammals then we wouldpredict that the single dose used by Sneddon would be aneffective antinociceptive for more than 16days. Furthermore, ifthe lag in antinociception relative to concentration in trout issimilar to that reported for mammals, then the effective timewould double to 32days in the studies of Sneddon from thesingle dose.

Our study was prompted by Jansen and Greene's report(1970) of the extremely rapid movement of morphine acrossfish gills. Our results do not support their contention of rapidtransport across the gills and further imply that adding morphineto the water is not an appropriate administration route. Further,our results show that the pharmacokinetics of morphine in fishis similar to mammals in that it shows a bi-exponential decreasein concentration after a single injection. The kinetic parametersdiffer from those in mammals in that the elimination phase is

about one order of magnitude slower in fish. Further research infish is needed to understand the distribution of the drug, howand where it is metabolized, what the metabolites are, theefficacy of the metabolites as analgesics, and how themetabolites are excreted. It also would be important to knowthe concentration of morphine in the brain and spinal cordrelative to the plasma concentration.

Acknowledgments

We thank Dr. Atef Mansour for the use of his spectropho-tometer, Dr. Bob Balahura for his valuable insights regardingmorphine chemistry, Dr. Craig Mosley for his insights regardingthe use of analgesics in comparative medicine, Dr. JanaSawchuck for her insights regarding pain studies in mammals,Dr. Mike Wilkie for his help with the ELISA, and Danny Boyce(ARDF, MUN) for providing the facilities which permittedthese studies. We would also like to thank Dr. Gilly Griffin fromthe Canadian Council of Animal Care for continued moralsupport and Dr. Chris Harvey-Clarke who convinced EDS toparticipate in the writing of the CCAC fish guidelines thatprompted this study. These studies were funded by NaturalSciences and Engineering Research Council of Canada(NSERC) discovery grants to EDS and AKG, by a Foundationfor Science and Technology doctoral fellowship (Portugal) toPCM, and by funds made available to the OSC throughNSERC's major facilities access program.

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