Measurement of peroxisomal enzyme activities in the liver of brown trout (Salmo trutta), using spectrophotometric methods
© Rocha et al; licensee BioMed Central Ltd. 2003
Received: 7 September 2002
Accepted: 11 March 2003
Published: 11 March 2003
This study was aimed primarily at testing in the liver of brown trout (Salmo trutta) spectrophotometric methods previously used to measure the activities of catalase and hydrogen peroxide producing oxidases in mammals. To evaluate the influence of temperature on the activities of those peroxisomal enzymes was the second objective. A third goal of this work was the study of enzyme distribution in crude cell fractions of brown trout liver.
The assays revealed a linear increase in the activity of all peroxisomal enzymes as the temperature rose from 10° to 37°C. However, while the activities of hydrogen peroxide producing oxidases were strongly influenced by temperature, catalase activity was only slightly affected. A crude fraction enriched with peroxisomes was obtained by differential centrifugation of liver homogenates, and the contamination by other organelles was evaluated by the activities of marker enzymes for mitochondria (succinate dehydrogenase), lysosomes (aryl sulphatase) and microsomes (NADPH cytochrome c reductase). For peroxisomal enzymes, the activities per mg of protein (specific activity) in liver homogenates were strongly correlated with the activities per g of liver and with the total activities per liver. These correlations were not obtained with crude peroxisomal fractions.
The spectrophotometric protocols originally used to quantify the activity of mammalian peroxisomal enzymes can be successfully applied to the study of those enzymes in brown trout. Because the activity of all studied peroxisomal enzymes rose in a linear mode with temperature, their activities can be correctly measured between 10° and 37°C. Probably due to contamination by other organelles and losses of soluble matrix enzymes during homogenisation, enzyme activities in crude peroxisomal fractions do not correlate with the activities in liver homogenates. Thus, total homogenates will be used in future seasonal and toxicological studies of brown trout peroxisomes.
A literature survey on animal peroxisomes reveals that most studies are confined to laboratory mammals and humans. The scarcity of knowledge on other vertebrate and invertebrate species, and the need for more comparative studies has been recently emphasised [1–3]. Nevertheless, it is known that fish peroxisomes have enzymatic systems similar to those reported in mammals. Catalase, D-amino acid oxidase, urate oxidase, allantoinase, allantoicase, L-α-hydroxyacid oxidase, alanine:glyoxylate aminotransferase, fatty acyl-CoA oxidase, enoyl-CoA hydratase and carnitine acetyltransferase are among the enzymes that were detected in fish peroxisomes [3–10]. It is also known that peroxisomal metabolism is lower in fish than in mammals. For example, in the liver of rainbow trout (Onchorynchus mykiss) and Japanese medaka (Oryzias latipes) peroxisomal enzyme activities are at least four folds lower than in rodent liver. These results were either assigned to a reduced number of peroxisomes per hepatocyte, or a lower amount of peroxisomal enzymes, or both .
The research in fish revealed that environmental factors like water temperature, salinity, season and feeding habits, exerted changes in peroxisomal enzyme activities that, additionally, vary greatly among species . It was also discovered that season, age and gender affect the morphology of fish liver peroxisomes. In the gray mullet (Mugil cephalus) hepatic peroxisomes tend to be larger in summer and in aged animals . Moreover, in the liver of brown trout (Salmo trutta) both the individual dimensions of peroxisomes and their total volume per hepatocyte, but not their number, change during the annual breeding cycle in both genders . Yet, in the latter species hepatic peroxisomes are significantly smaller in females than in males during the breeding season . These observations raised the question about the kind of biochemical changes that may be associated with those morphological variations detected in fish liver peroxisomes. To measure some peroxisomal enzyme activities in brown trout liver, a method originally applied for mammalian peroxisomal hydrogen peroxide producing oxidases  was tested for the first time in fish. Catalase, a well-known peroxisomal marker enzyme , was also included in this work. Since the temperature is an important factor that affects enzyme activities, its influence on several trout peroxisomal enzymes was investigated. This work also aimed at studying the distribution of several peroxisomal enzymes in crude cell fractions. To control the amount of cross contamination in the crude peroxisomal fraction by other organelles, standard spectrophotometric procedures were used in order to measure the activity of marker enzymes of mitochondria (succinate dehydrogenase), lysosomes (aryl sulphatase) and microsomes (NADPH cytochrome c reductase). Finally, this paper reports the first data about the activities of peroxisomal enzymes in the liver of brown trout.
Temperature and sample concentration tests
Since protein content and enzyme activities in brown trout liver were much different from those reported in rodent liver, several sample dilutions were tested in order to determine the best proportion between protein concentration and the concentration of all other components in the incubation medium. Therefore, after the dilution of homogenates and fractions the following final protein concentrations were used: 2 and 4 μg ml-1 for catalase, 45 and 90 μg ml-1 for D-aminoacid oxidase and fatty acyl-CoA oxidase, 10 and 20 μg ml-1 for urate oxidase, 0.15 and 0.3 mg ml-1 for glycolate oxidase and succinate dehydrogenase, 0.2 and 0.4 mg ml-1 for both aryl sulphatase and NADPH cytochrome c reductase. With these two dilutions for each enzyme, activities were linear in time and a good proportionality between enzymatic activity and the amount of protein was obtained. These sample concentrations also provided results with low variability between the two dilutions used (Coefficient of Variation ≤ 0.05) and high reproducibility between replicates of the same dilution (CV ≤ 0.02).
Measurement of enzyme activities
Specific and per g enzyme activities in brown trout liver
Activity per g of liver
s-1 g-1 liver (*)
or nmol min-1 g-1 liver
s-1 mg-1 protein (*) or
nmol min-1 mg-1 protein
Fatty acyl-CoA oxidase
NADPH cytochrome c reductase
Protein per g of liver
(mg g-1 liver)
Total protein per fraction
Percentage distribution of the marker enzymes
Percentage Distribution (%)
Fatty acyl-CoA oxidase
NADPH cytochrome c reductase
Fraction D also showed high percentage distributions (Table 2) and elevated specific activities for the enzymatic markers of both mitochondria (succinate dehydrogenase) and lysosomes (aryl sulphatase) (Table 1). Moreover, the relative specific activities of succinate dehydrogenase and aryl sulphatase showed that they were more or less equally distributed both in B and D fractions (Fig. 2). Nonetheless, percentage distribution, specific activity and relative specific activities of these two enzymes were lower in E and F fractions than in the other fractions (Fig. 2, Tables 1 and 2). Considering the microsomal marker enzyme NADPH cytochrome c reductase, it was observed that in D fraction its activity was less than half of that determined in E fraction (Table 1).
The percentage distribution of peroxisomal enzymes in D fraction (Table 2) ranged from 22.7 for fatty acyl-CoA oxidase to 34.2 for catalase, but according to ANOVA such differences have no statistical significance. The same was verified in both B and E fractions. However, in F fraction the percentage distribution of catalase (38.2) is significantly different from the percentage distribution of fatty acyl-CoA oxidase (66.5), although no other statistical differences among peroxisomal enzymes were found within this fraction (Table 2).
Statistical analyses also revealed that specific activities of peroxisomal enzymes in D fraction did not have any explicit pattern of correlation with the specific activities in fraction A, neither with the activities per g of liver or per liver. Contrastingly, in fraction A it was observed that for each peroxisomal enzyme there was a strong and positive correlation between the specific activity and the activity per g of liver: urate oxidase (r = 0.96; p = 0.002); D-aminoacid oxidase (r = 0.99; p = 0.0002); fatty acyl-CoA oxidase (r = 0.96; p = 0.002); glycolate oxidase (r = 0.95; p = 0.003); catalase (r = 0.89; p = 0.02). In this fraction, the specific activity of each oxidase was also highly correlated with the total activity per liver: urate oxidase (r = 0.97; p = 0.001); D-aminoacid oxidase (r = 0.86; p = 0.03); fatty acyl-CoA oxidase (r = 0.97; p = 0.001); glycolate oxidase (r = 0.94; p = 0.005).
One of the purposes of this study was to test in brown trout liver the spectrophotometric method originally used by Cablé et al.  for measurement of peroxisomal oxidase activities in mammalian tissues. However, some modifications were introduced and the method was extended to urate and glycolate oxidases. To calculate the amounts of H2O2 produced by these enzymes, instead of using the extinction coefficient previously published , a calibration line was made with known amounts of H2O2. Others also made similar calibration lines, but when using luminometric methods  and fluorometric techniques .
Since the physiological temperature is much lower in brown trout than in mammals, temperature influence in the activity of five peroxisomal enzymes was investigated. The results showed that the activities of these enzymes rose in a linear mode from 10 to 37°C, but while the activity of hydrogen peroxide producing oxidases were strongly influenced by temperature, catalase activity was only slight affected. This last observation is in total accordance with previous data regarding catalase activity in mammals . Therefore, these enzymatic activities can be measured at whatever temperature within the above-referred range. However, at 37°C activities are higher and therefore easier to measure than at the physiological temperatures of brown trout (10 – 15°C). In fact, other authors have used a wide range of temperatures in studies involving fish peroxisomal enzymes. For example, in carp (Cyprinus carpio) catalase activity was determined at 25°C whereas urate oxidase, fatty acyl-CoA oxidase and glycolate oxidase were measured at 37°C [4, 5, 17] and in rainbow trout (O. mykiss) catalase activity was determined on ice whereas fatty acyl-CoA oxidase was measured at 15°C .
Some similarities are evident when crude cell fractions of rat liver  are compared with those obtained from brown trout liver. In both species relative specific activities of catalase and hydrogen peroxide producing oxidases are higher in the D fraction obtained using very similar centrifugation procedures. Moreover, catalase and glycolate oxidase (or L-α-hydroxyacid oxidase) present a high percentage distribution in the cytosolic fraction of rat and brown trout, has expected for soluble matrix enzymes released from peroxisomes due to membrane damage during homogenisation. On the other hand, urate oxidase is not detectable in the supernatant of rat liver  while in brown trout about 50% of urate oxidase activity can be found in the supernatant. This discrepancy is due to a fundamental difference between brown trout and mammalian urate oxidase, which behaves exactly like a soluble matrix enzyme in the former but is a crystallised core enzyme in the latter. Indeed cores were neither found in brown trout liver peroxisomes  or in other fish species . Although urate oxidase was described as a membrane-linked enzyme in carp liver peroxisomes  in other fish species was considered a soluble matrix enzyme  as it seems to be in brown trout liver peroxisomes.
Differences between rat and brown trout crude liver fractions are also evident in the microsomal fraction, in which the percentage distributions of peroxisomal enzyme activities are higher in rat than in brown trout .
The absence of correlations between D fractions and total liver homogenates showed that enzyme activities in peroxisomal enriched fractions are not proportional to the activities in homogenates. The contamination of crude peroxisomal fractions with other organelles and the substantial loss of soluble matrix enzymes can explain this result. Because of these facts, and knowing that in rainbow trout the gradient centrifugation of the crude peroxisomal fraction produced a further decrease in the activities of some oxidases , in future seasonal and toxicological studies about brown trout peroxisomes total homogenates will be used instead of cell fractions. Moreover, variation coefficients suggest that enzyme activities have relevant individual variability in brown trout. Thus, very subtle differences due to experiments or natural events may pass unnoticed when working with a small number of animals.
In conclusion: 1) the spectrophotometric protocols used for measurement of mammalian peroxisomal enzymes can be successfully applied to brown trout; 2) the enzymatic activities of the peroxisomal enzymes studied by us can be correctly measured between 10° and 37°C, this because their activity rose in a linear mode with temperature, with catalase activity increasing only slightly; 3) for peroxisomal enzymes in brown trout liver homogenates there is a clear correlation between specific activity and activity per g of liver or per liver; 4) during preparation of cell fractions enriched with peroxisomes a significant amount of matrix enzymes are loosed, and the remaining enzyme activities are not correlated with the activities in liver homogenates.
Two-year-old brown trout males (n = 6) were collected by random net fishing from aquaculture pools (Posto Aquícola do Torno – Amarante, Portugal) in late February. Fish of 188 g (coefficient of variation, CV = 0.10) in weight and 26 cm (CV = 0.04) in length were kept in dechlorinated water (pH 6–7, at 8–10°C) for 1–2 days before sacrifice. Mean liver weight was 3.0 g (CV = 0.30).
Cofactors, substrates for enzyme assays and bovine serum albumin (BSA) were purchased from Sigma Chemical Co. (Poole, Dorset, U.K). Other chemicals were obtained from Merck (Darmstadt, Germany).
Animals were anaesthetised by bathing in a solution of ethylene glycol monophenyl ether (0.4 ml l-1) and then perfused at 5.6 ml min-1 kg-1 via the portal venous system for 4–5 min with an isosmotic buffer for salmonids . The liver was weighted just after removal, immediately minced in chilled homogenization buffer with a pH of 7.4 (250 mM sucrose, 5 mM MOPS, 1 mM EDTA and 0.1% ethanol saturated with phenyl-methyl-sulfonyl-fluoride PMSF)  and then homogenized in the same buffer using a Potter-Elvehjem homogenizer at 1000 rpm, held in an ice-water-bath. After sedimentation of unhomogenized material (50 g, 10 min, 4°C) according to Cajaraville et al. , supernatant volume was adjusted to 5 ml g-1 of liver using ice-cold homogenisation buffer (A fraction). A previously described method  was used for differential pelleting of the A fraction in order to produce a B fraction containing mainly mitochondria and lysosomes (2,000 g, 10 min, 4°C), a D fraction enriched with peroxisomes (20,000 g, 30 min, 4°C), a E fraction enriched with microsomes (100,000 g, 60 min, 4°C) and a cytosolic F fraction (supernatant of 100,000 g, 60 min, 4°C). Pellets (B, D and E) were rinsed once by resuspension in an appropriate volume of buffer using a glass-rod and recentrifuged under the same conditions. Before enzymatic assay, all fractions were treated with Triton X-100 at a final concentration of 0.5% (v/v) in homogenization buffer. Proteins were assayed according to Lowry et al.  using BSA standards and results are expressed in BSA equivalents (Table 1).
Enzymatic assays were carried out in a spectrophotometer connected to a circulating water system for temperature regulation in the cuvette compartment. Enzyme activities linear in time and proportional to the amount protein in the assays were obtained using an appropriate sample dilution. Two different dilutions of each sample were used to produce a mean value of enzymatic activities. Temperature effect on peroxisomal enzyme activities was evaluated by measurements made at 10, 15, 20, 25, 30 and 37°C.
For catalase the methodology previously described by Aebi  was used. Incubation medium contained 50 mM sodium phosphate buffer (pH 7), 10 mM hydrogen peroxide (H2O2) and a diluted sample. Consumption of H2O2 was followed by the decrease in absorbance at 240 nm for 30 seconds and the activity is expressed by the first-order rate constant (k) for degradation of H2O2. k = (1 ÷ Δt) × ln (c1 ÷ c2) where c1 and c2 correspond to H2O2 concentrations at t = 0 and t = 30 seconds, respectively .
Measurement of peroxisomal oxidase activities was based on the production of H2O2 and followed the procedure of Cablé et al.  with some modifications. Incubation medium contained 50 mM potassium phosphate buffer (pH 8.3), 0.082 mM 4-amino-antipyrine, 1 mM phenol and 2 IU ml-1 of horseradish peroxidase. To avoid the interference of catalase, 10 mM of azide was added to the medium, according to Leupold et al. . For glycolate oxidase assay, 0.01 mM flavin mononucleotide (FMN) was added in the medium . For other oxidases, 0.01 mM flavin adenine dinucleotide (FAD) was included in the medium [13, 23]. Substrates were used in the following concentrations: 20 mM D-alanine, 1 mM uric acid, 0.1 mM palmitoyl-CoA (C:16) and 20 mM sodium glycolate. Each enzymatic reaction was started by addition of 25 μl of a diluted sample to 650 μl of incubation medium. Absorbance increase was measured at 500 nm for 10 min. For urate oxidase, a baseline was made with the complete medium without sample, which was thereafter subtracted from the absorbance increase in each assay. This procedure was not necessary for all other assayed oxidases, because non-specific reactions were not detected in other cases.
Succinate dehydrogenase activity was estimated according to Schoner et al. . Incubation medium contained 50 mM potassium phosphate buffer (pH 7.5), 5 mM sodium succinate, 1 mM potassium cyanide, 0.1 mM 2,6-dichlorophenol indophenol (DPIP), and a diluted sample. The extinction coefficient of DPIP at 600 nm, determined with several standard solutions ranging from 0 to 100 μM of DPIP (ε = 17.3 mM-1 cm-1), was used to calculate the activity .
Measurements of aryl sulphatase activity followed the method of Worwood et al. . Incubation medium contained 0.5 M acetate buffer (pH 5.6), 20 mM nitrocatechol sulphate and a diluted sample. The reaction was stopped by addition 1 ml of 1 M sodium hydroxide and the extinction coefficient of nitrocatechol solutions at 515 nm (ε = 12.4 mM-1 cm-1) was used to calculate the activity .
NADPH cytochrome c reductase
Measurements of NADPH cytochrome c reductase activity followed the method of Johannesen et al. . Incubation medium contained 0.1 M TRIS-HCl buffer (pH 7.6), 10 mM potassium cyanide, 0.4 mM oxidised cytochrome c, 3 mM NADPH and a diluted sample. This reaction was started by addition of NADPH. Activity calculations were based upon the extinction coefficient of a reduced cytochrome c solution at 550 nm (ε = 19.6 mM-1 cm-1) .
Calculations, units and statistical analysis
The percentage distribution [(protein or enzyme activity in one fraction ÷ Σ of protein or enzyme activity in fractions B to F) × 100] and the relative specific activity [percentage distribution of the enzyme in one fraction ÷ percentage distribution of protein in that fraction] were calculated as in other papers . Catalase activity is expressed by the first-order rate constant in s-1 g-1 of liver and s-1 mg-1 of protein, for all others, results are given in nmol min-1 g-1 of liver and nmol min-1 mg-1 of protein.
Data are reported as means per group of animals, followed by the respective coefficients of variations (CV = standard deviation ÷ mean). Correlation analysis was used to establish several relationships considered biologically or technically relevant. Results were judged significant when p ≤ 0.05 as reported by Pearson's coefficient of correlation (r). A linear regression analysis was made to study enzyme activities at six different chosen temperatures.
The fish were generously provided by the "Divisão de Caça e Pesca nas Águas Interiores – Direcção Regional de Agricultura de Entre-Douro e Minho", under an official cooperation protocol. The commitment of Eng. Álvaro Gonçalves and Eng. Augusto Maia is particularly appreciated. This study was financially supported by the pluriannual program of the CIIMAR and by the "Fundação para a Ciência e a Tecnologia" (research contract n° PRAXIS/C/BIA/14268/1998).
- Fahimi HD, Cajaraville MP: Induction of peroxisome proliferation by some environmental pollutants and chemicals in animal tissues. In: Cell Biology in Environmental Toxicology. Edited by: Cajaraville MP. 1995, Basque Country Press Service, Bilbo, 221-255.Google Scholar
- Lobo-da-Cunha A: The peroxisomes of the hepatopancreas in two species of chitons. Cell Tissue Res. 1977, 290: 655-664. 10.1007/s004410050971.View ArticleGoogle Scholar
- Cancio I, Cajaraville MP: Cell biology of peroxisomes and their characteristics in aquatic organisms. Int Rev Cytol. 2000, 199: 201-293.View ArticlePubMedGoogle Scholar
- Kramar R, Goldenberg H, Böck P, Klobucar N: Peroxisomes in the liver of the carp (Cyprinus carpio L.) electron microscopic cytochemical and biochemical studies. Histochem. 1974, 40: 137-154.View ArticleGoogle Scholar
- Goldenberg H, Hüttinger M, Kampfer P, Kramar R: Preparation of peroxisomes from carp liver by zonal rotor density gradient centrifugation. Histochem J. 1978, 10: 103-113.View ArticlePubMedGoogle Scholar
- Noguchi T, Takada Y, Fujiwara S: Degradation of uric acid to urea and glyoxylate in peroxisomes. J Biol Chem. 1979, 254: 5272-5275.PubMedGoogle Scholar
- Baldwin LA, Calabrese EJ, Kostecki PT, Yang J-H: Isolation of peroxisomal enoyl-CoA hydratase in rainbow trout and immunochemical identification with the bifuntional enzyme. Fish Physiol Biochem. 1990, 8: 347-351.View ArticlePubMedGoogle Scholar
- Braunbeck T, Völkl A: Induction of biotransformation in the liver of eel (Anguilla anguilla) by sublethal exposure to dinitro-o-kresol: an ultrastructural and biochemical study. Ecotoxicol Environ Saf. 1991, 21: 109-127.View ArticlePubMedGoogle Scholar
- Scarano LJ, Calabrese EJ, Kostecki PT, Baldwin LA, Leonard DA: Evaluation of a rodent peroxisome proliferator in two species of freshwater fish: rainbow trout (Onchorynchus mykiss) and Japanese medaka (Oryzias latipes). Ecotoxicol Environ Saf. 1994, 29: 13-19.View ArticlePubMedGoogle Scholar
- Sakuraba H, Fujiwara S, Noguchi T: Metabolism of glyoxylate, the en product of purine degradation in liver peroxisomes of fresh water fish. Biochem Biophys Res Commun. 1996, 229: 603-606. 10.1006/bbrc.1996.1850.View ArticlePubMedGoogle Scholar
- Orbea A, Beier K, Völkl A, Fahimi HD, Cajaraville MP: Ultrastructural, immunocytochemical and morphometric characterisation of liver peroxisomes in gray mullet, Mugil cephalus. Cell Tissue Res. 1999, 297: 493-502. 10.1007/s004410051376.View ArticlePubMedGoogle Scholar
- Rocha E, Lobo-da-Cunha A, Monteiro RAF, Silva MW, Oliveira MH: A stereological study along the year on the hepatocytic peroxisomes of brown trout (Salmo trutta). J Submicr Cytol Pathol. 1999, 31: 91-105.Google Scholar
- Cablé S, Kedinger M, Dauça M: Peroxisomes and peroxisomal enzymes along the crypt-villus axis of the rat intestine. Differentiation. 1993, 54: 99-108.View ArticlePubMedGoogle Scholar
- Aebi H: Catalase in vitro. Method Enzymol. 1984, 105: 121-126.View ArticleGoogle Scholar
- Leupold C, Völkl A, Fahimi HD: Luminometric determination of oxidase activity in peroxisomal fractions of rat liver: Glycolate oxidase. Anal Biochem. 1985, 151: 63-69.View ArticlePubMedGoogle Scholar
- Walusimbi-Kisitu M, Harrison EH: Fluorometric assay for rat liver peroxisomal fatty acyl-coenzyme A oxidase activity. J Lipid Res. 1983, 24: 1077-1084.PubMedGoogle Scholar
- Goldenberg H: Organisation of purine degradation in the liver of a teleost (carp; Cyprinus carpio L.). A study of its subcellular distribution. Molec Cell Biochem. 1977, 16: 17-21.View ArticlePubMedGoogle Scholar
- Henderson RJ, Sargent JR: Chain-length specificities of mitochondrial and peroxisomal β-oxidation of fatty acids in livers of rainbow trout (Salmo gardneri). Comp Biochem Physiol. 1985, 82: 79-85. 10.1016/0305-0491(85)90131-2.Google Scholar
- Völkl AE, Fahimi HD: Isolation and characterisation of peroxisomes from the liver of normal untreated rats. Eur J Biochem. 1985, 149: 257-265.View ArticlePubMedGoogle Scholar
- Rocha E, Monteiro RAF, Pereira CA: The liver of the brown trout, Salmo trutta fario: a light and electron microscope study. J Anat. 1994, 185: 241-249.PubMed CentralPubMedGoogle Scholar
- Cajaraville MP, Völkl A, Fahimi HD: Peroxisomes in digestive gland cells of the mussel Mytilus galloprovincialis Lmk. Biochemical, ultrastructural and immunocytochemical characterisation. Eur J Cell Biol. 1992, 59: 255-264.PubMedGoogle Scholar
- Lowry OH, Rosebrough NJ, Farr AL, Randall RJ: Protein measurement with the Folin phenol reagent. J Biol Chem. 1951, 193: 265-275.PubMedGoogle Scholar
- Vamecq J: Fluorometric assay of peroxisomal oxidases. Anal Biochem. 1990, 186: 340-349.View ArticlePubMedGoogle Scholar
- Nelson DP, Kiesow LA: Enthalpy of decomposition of hydrogen peroxide by catalase at 25°C (with molar extinction coefficients of H2O2 solutions in the UV). Anal Biochem. 1972, 49: 472-478.View ArticleGoogle Scholar
- Schoner W, von Ilberg C, Kramer R, Seubert W: On the mechanism of Na+- and K+-stimulated hydrolysis of adenosine triphosphate. 1. Purification and properties of a Na+- and K+-activated ATPase from ox brain. Euro J Biochem. 1967, 1: 334-343.View ArticleGoogle Scholar
- Worwood M, Dodgson KS, Hook GER, Rose FA: Problems associated with the assay of arylsulphatases A and B of rat tissues. Biochem J. 1973, 134: 183-190.PubMed CentralView ArticlePubMedGoogle Scholar
- Johannesen K, Depierre JW, Bergstrand A, Dallner G, Ernster L: Preparation and characterisation of total, rough and smooth microsomes from the lung of control and methylcholanthrene-treated rats. Biochem Biophys Acta. 1977, 496: 115-135. 10.1016/0304-4165(77)90120-9.View ArticlePubMedGoogle Scholar
- Livingstone DR, Moore MN, Widdows J: Ecotoxicology: biological effects measurements on molluscs and their use in impact assessment. In: Pollution of the North Sea: an assessment. Edited by: Salomons W, Duvrsma EK, Bayne BL, Forstner U. 1988, Spriger-Verlag, Berlin, Heidelberg, New York, 624-637.Google Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article: verbatim copying and redistribution of this article are permitted in all media for any purpose, provided this notice is preserved along with the article's original URL.