Metabolism

Metabolism T. Ikeda, JJ. Torres, S. Hernandez-Leon and S.P. Geiger Review 1 10.1 OXYGEN CONSUMPTION AS AN INDEX OF METABOLISM 'Metabolism' may be defined as the sum of all processes through which protoplasm is formed from food (anabolism) and broken down into waste matter (catabolism), with release of energy. Energy gained through the processes is stored exclusively in the form of adenosine triphosphate (ATP), and used for various activities (locomotion, produc tion of new tissue, ion pumps etc.) of living organisms. ATP is generated from the tricarboxylic-acid (TCA) cycle and oxygen is required to drive the cycle. Therefore metabolic rate may be estimated by measuring oxygen consumption rate. Exceptions are anaerobiosis in intertidal organisms and burst escape activity, but the oxygen debt incurred is usually repaid later when oxygen is once more available (cf. Clarke 1987).'External respiration' is the mechanism by which oxygen is brought into and carbon dioxide expelled from the organism, whereas 'internal respiration' refers to the sum of enzymatic reactions, both oxidative and nonoxidative, by which energy is made available for biological work (Prosser 1961). In most ecological studies dealing with intact organisms, 'external respiration' is of special interest and is simply referred as 'respiration'. For this reason, oxygen consumption rate is referred to as respiration (or respiratory) rate or vice versa. In this sense, either oxygen consumption or carbon dioxide production can be used as an index of metabolic rate of marine zooplankton. However, accurate determination of carbon dioxide in sea water is hampered by the strong buffering action of sea water and has been used in only a few studies, i.e. Neomysis integer by Raymont and Krishnaswamy (1968) and Rhincalanus gigas by Rakusa Suszczewski et al. (1976). In contrast, accurate determination of dissolved oxygen in sea water became possible with the advent of the Winkler titration method in the late 1800s (Winkler 1888). In the early 1960s, oxygen-electrodes were introduced to the study of marine zooplankton metabolism as a simple tool for measuring dissolved oxygen in sea water (Teal and Halcrow 1962; Halcrow 1963). Examples of very early studies on the oxygen consumption rates of marine zooplank ton may be found in the work of Vernon (1896) on salps, followed by that of Ostenfield (1913, cited in Marshall 1973) on the copepod Calanus hyperboreus. Since then voluminous data have been accumulated on the metabolism of various zooplankton species living in many regions of the world's oceans (for review, see Marshall 1 973; Ikeda 1974; Conover 1978; Raymont 1983). Part of the data on selected zooplankton species are shown in Table 10.1. Omori and Ikeda (1984) and Le Borgne (1986) reviewed methodologies for studies of marine zooplankton metabolism. The metabolic rate of animals is defined with respect to the activity of animals as follows: 'standard' (or 'basal') metabolism is the oxygen consumption rate for mainICES Zooplankton Methodology Manual Copyright @ 2000 Academic Press ISBN 01 2-327645-4 All rights of reproduction in any form reserved Key words: Zooplankton, Respiration, Excretion, ETS, GDH 10 Metabolism T. Ikeda, J.J. Torres, S. Hernández-León and S.P. Geiger Review 110.1 OXYGEN CONSUMPTION AS AN INDEX OF METABOLlSM 'Metabolism' may be defined as the sum of aH processes through which protoplasm is formed from food (anabolism) and broken down into waste matter (catabolism), with release of energy. Energy gained through the processes is stored exclusively in the form of adenosine triphosphate (ATP), and used for various activities (locomotion, production of new tissue, ion pumps etc.) of living organisms. ATP is generated from the tricarboxylic-acid (TCA) cycle and oxygen is required to drive the cycle. Therefore metabolic rate may be estimated by measuring oxygen consumption rateo Exceptions are anaerobio si s in intertidal organisms and burst escape activity, but the oxygen debt incurred is usually repaid later when oxygen is once more available (cf. Clarke 1987). 'External respiration' is the mechanism by which oxygen is brought into and carbon dioxide expelled from the organism, whereas 'internal respiration' refers to the sum of enzymatic reactions, both oxidative and nonoxidative, by which energy is made available for biological work (Prosser 1961). In most ecological studies dealing with intact organisms, 'external respiration' is of special interest and is simply referred as 'respiration'. For this reason, oxygen consumption rate is referred to as respiration (or respiratory) rate or vice versa. In this sense, either oxygen consumption or carbon dioxide production can be used as an index of metabolic rate of marine zooplankton. However, accurate determination of carbon dioxide in sea water is hampered by the strong buffering action of sea water and has been used in only a few studies, i.e. Neomysis integer by Raymont and Krishnaswamy (1968) and Rhincalanus gigas by RakusaSuszczewski et al. (1976). In contrast, accurate determination of dissolved oxygen in sea water became possible with the advent of the Winkler titration method in the late 1800s (Winkler 1888). In the early 1960s, oxygen-electrodes were introduced to the study of marine zooplankton metabolism as a simple tool for measuring dissolved oxygen in sea water (Te al and Halcrow 1962; Halcrow 1963). Examples of very early studies on the oxygen consumption rates of marine zooplankton may be found in the work of Vernon (1896) on salps, followed by that of Ostenfield (1913, cited in Marshall 1973) on the copepod Calanus hyperboreus. Since then voluminous data have been accumulated on the metabolism of various zooplankton species living in many regions ofthe world's oceans (for review, see Marshall1973; Ikeda 1974; Conover 1978; Raymont 1983). Part of the data on selected zooplankton species are shown in Table 10.1. Omori and Ikeda (1984) and Le Borgne (1986) reviewed methodologies for studies of marine zooplankton metabolismo The metabolic rate of animal s is defined with respect to the activity of animal s as foHows: 'standard' (or 'basal') metabolism is the oxygen consumption rate for mainICES Zooplankton Methodology Manual ISBN 0-12-327645-4 Copyright © 2000 Academic Press AH rights of reproduction in any fonn reserved Table 10.1 Rates of oxygen consumption, ammonia excretion and inorganic phosphate excretion of various marine zooplankton species determined by incubating specimens in filtered sea water.-= no data, indiv = individual. Ex perim enta l O xygen A m m o nia Ph osp hate tem pe rature D W co n su m ption ex cretio n excretio n Zo op lan kto nspecie s Lo cation (C) (m g) (/ilO 2ind iv 1 h  1) O zgN in div  1h  1) (^ g P ind iv l h " 1) So u rce Co ele nterata A g lan thad ig ita te Ba re ntsSea 1.1 14.0 2 .27 0 .082 0.035 Ike daandSkio ld al(19 89) Cte no ph ora M n em iopsisleidyi N arrag ansett 20 299 30 .35 2 .93 1.00 K rem er(197 7) Bay Thecosom ata Ca vo lin ialong irostris G B Rinsh o re 27 8.9 16 .7 1.8 1 0.30 Iked aandSkio ld al w ater (un pu b l.) Lim a cinah elicina Bare ntsSea 0 .0 Bare ntsSea ‑ 0.1 0.593 0 .4 51 16.47 1.6 5 0 .0 072 0.0052 Iked aandSkio ld al(1989 ) G ym noso m ata Clion elim a cin a 0 .0 34 0.021 Iked aandSkio ld al(1989) Co pe pod a Ca lanu sfin m arch icus,C 69 BarentsSea 0 .1 0 .387 0.328 0 .0 13 0.0050 Iked aandSkio lda l(1989) C.g lacialis,C 6$ BarentsSea 3 .5 0 .410 0.630 0 .0 26 0 .010 Iked aandSkio lda l(1989) C.hyp erb oreus,C 6? BarentsSea 1.3 3.95 1.49 0.049 0 .0050 Iked aa ndSkio lda l(1989) N eo calan uscrista tus,C 5 o ffH o kkaid o 6.3 1.59 1.67 0.164 ‑ 1ked a(1974 ) N .p lum ch rus,C 5 o ffH o kkaid o 7.3 0 .785 0.68 0.0 17 ‑ 1ked a(1974) Eucalan usbu ng ii,C6$ o ffH ok kaid o 6.0 1.01 0.80 0.0 58 ‑ 1ked a(1974) M etrid iap acifica,C6$ o ffH ok kaid o 7.7 0 .182 0.383 0.0 19 ‑ 1ked a(1974) A cartiaa ustralis G BRinsho re 24.5 0 .009 1 0.056 0.00 54 0 .0 018 Iked aa ndSkio lda l(1980) w ater U n din ulad arw in i Ind ianO cean 28.5 0 .15 1.13 0.137 0 .044 G au dyandBo uch er(1983) Euchae tam arina Ind ianO cean 28.5 0 .22 1.30 0.13 1 0 .0 52 G au dyandBo uch er(1983) Tem orad iscaud ata Ind ianO cean 28.5 0 .034 0.23 0.020 0 .0 10 G au dyandBo uch er(1983) Key words: Zooplankton, Respiration, Excretion, ETS, GDH Table 10.1 Rates of oxygen consumption, ammonia excretion and inorganic phosphate excretion of various marine zooplankton species determined by incubating specimens in filtered sea water. -= no data, indiv = individual. Experimental Oxygen Ammonia Phosphate temperature DW consumption excretion excretion Zooplankton species Location eC) (mg) (pi O2 indiv1 h1) (JlgN indiv1 h1) (JlgP indiv1 h1) Source Coelenterata Aglantha digitale Barents Sea 1.1 14.0 2.27 0.082 0.035 Ikeda and Skjoldal (1989) Ctenophora Mnemiopsis leidyi Narragansett 20 299 30.35 2.93 1.00 Kremer (1977) Bay Thecosomata Cavolinia longirostris GBR inshore 27 8.9 16.7 1.81 0.30 Ikeda and Skjoldal water (unpubl.) Limacina helicina Barents Sea 0.0 0.593 0.451 0.0072 0.0052 Ikeda and Skjoldal (1989) Gymnosomata Clione limacina Barents Sea -0.1 16.47 1.65 0.034 0.021 Ikeda and Skjoldal (1989) Cope poda Calanus finmarchicus, C6~ Barents Sea 0.1 0.387 0.328 0.013 0.0050 Ikeda and Skjoldal (1989) C. glacialis, C6~ Barents Sea 3.5 0.410 0.630 0.026 0.010 Ikeda and Skjoldal (1989) C.hyperboreu~C6~ Barents Sea 1.3 3.95 1.49 0.049 0.0050 Ikeda and Skjoldal (1989) Neocalanus cristatus, C5 off Hokkaido 6.3 1.59 1.67 0.164 Ikeda (1974) N. plumchrus, C5 off Hokkaido 7.3 0.785 0.68 0.017 Ikeda (1974) Eucalanus bungii, C6~ off Hokkaido 6.0 1.01 0.80 0.058 Ikeda (1974) Metridia pacifica, C6~ off Hokkaido 7.7 0.182 0.383 0.019 Ikeda (1974) Acartia australis GBR inshore 24.5 0.0091 0.056 0.0054 0.0018 Ikeda and Skjoldal (1980) water Undinula darwini Indian Ocean 28.5 0.15 1.13 0.137 0.044 Gaudy and Boucher (1983) Euchaeta marina Indian Ocean 28.5 0.22 1.30 0.131 0.052 Gaudy and Boucher (1983) Temora discaudata Indian Ocean 28.5 0.034 0.23 0.020 0.010 Gaudy and Boucher (1983) Mysidacea Acanthomysis pseudomacropsisCj. off Hokkaido 14.9 2.88 4.36 0.630 Ikeda (1974) Amphipoda Themisto libel/ula Barents Sea -0.1 2.98 2.53 0.053 0.018 Ikeda and Skjoldal (1989) Euphausiacea Thysanoessa inermis Barents Sea 1.9 34.78 13.77 0.855 0.33 Ikeda and Skjoldal (1989) Euphausia pacifica off Hokkaido 10.2 12.95 16.10 0.781 Ikeda (1974) E.lucens off SW Africa 12.5 5.00 10.47 0.850 Stuart (1986) Chaetognatha Sagitta elegans Barents Sea -0.3 4.50 1.41 0.345 0.028 Ikeda and Skjoldal (1989) S. enflata GBR inshore 25 0.71 1.67 0.12 0.021 Ikeda and Skjoldal water (unpubl.) Appendiculata Oikopleura dioica Lab culture 24 0.0032 0.209 0.0125 0.0012 Gorsky et al. (1987) Thaliacea Salpa thompsoni, sol. Antarctic -1.0 114.63 12.0 0.806 0.39 Ikeda and Mitchell (1982) water Thalia democratica, sol. off S. Africa 17.3 2.97 1.77 0.18 Ikeda (1974) 458 METABOLlSM tenance only, 'routine' metabolism is the oxygen consumption rate measured with uncontrolled but minimum motor activity, and 'active' metabolism is the oxygen consumption rate with enforced activity at a maximallevel. When the oxygen consumption rate is measured at different activity levels, the rate extrapolated to zero activity is the standard metabolismo In most studies on zooplankton metabolism, no attempts have been made to relate the oxygen consumption data to the level of activity of animal s on the premise that the measured rates are close to routine metabolismo As notable exceptions, Torres and Childress (1983) and Buskey (1998) established the relationship between oxygen consumption rates and swimming speed in the migrating euphausiid Euphausia pacifica and swarming copepod Dioithona oculata respectively. According to their results, standard metabolism is 0.7 and active metabolism is 2.7 times routine metabolism in E. pacifica, and respective figures are 0.5 and 3.3 times routine metabolism in D. oculata. As an alternative approach, net cost of swimming for several crustacean plankton has been calculated on the theoretical grounds of mechanical power dissipated as drag (cf. Morris et al. 1990, and references therein). 1 10.1.1 Conversion of oxygen consumption to carbon and calorific units The molar ratio of carbon dioxide produced to oxygen consumed is called the 'Respiratory Quotient' (RQ). In theory, the RQ varies from 0.71 to 1.0 depending on metabolic substrates (lipid, 0.71; protein, 0.80; carbohydrate, 1.0; from Prosser 1961). According to Gnaiger's (1983) re-calculation the RQ value for carbohydrate remains unchanged (1.0), but for lipid, it is 0.72 and for protein it changes as a function of the excretory end-product, i.e. 0.97 for ammonia and 0.84 for urea. Since marine zooplankton are considered to be primarily ammonotelic (see Nitrogen and phosphorous metabolism in section 10.2) an RQ of 0.97 may be more appropriate to convert their oxygen consumption data to carbon units; mi O2 (individual h)-t x RQ x 12/22.4 = mg e (individual h)-t (10.1) where 12/22.4 is the weight (12 g) of carbon in 1 mole of (22.4 1) of carbon dioxide. Oxygen consumption rate expressed in units of carbon represents the carbon requirement for zooplankton metabolism, and can be used as an index of minimum food requirements when assimilation efficiency and growth are not taken into account. The caloric equivalent to a unit volume of oxygen consumed varies depending on the metabolic substrate utilized. According to Gnaiger (1983), there are 440, 447 and 471 kilo joules (kJ) per mole of oxygen consumed for lipid, protein (ammonia as endproduct) and carbohydrate, respectively. Since 1 mole of oxygen is 22.41 and 1 calorie is equivalent to 4.1868 J, the oxycaloric equivalents can be expressed in calories as 4.69, 4.76 and 5.02 kcal per liter of oxygen consumed for lipid, protein and carbohydrate meta bolism, respectively. 1 10.2 NITROGEN ANO PHOSPHORUS METABOLlSM Animals produce various substances as end-products of metabolismo Although zooplankton excreta include both liquid and solid forms (fecal pellets) we will only consider he re liquid forms. Nitrogen compounds have been measured in terms of total N, ammonia-N, amino-N and urea-N, and phosphorus compounds in terms of total-P, inorganic-P and organic-P. Among these, ammonia and inorganic phosphorus are of NITROGEN AND PHOSPHORUS METABOLlSM 459 special interest because of their importance as immediately available nutrients for phytoplankton (cf. Corner and Davies 1971). Excretion rates of ammonia and inorganic phosphate ofselected zooplankton species are shown in Table 10.1. A major end-product of nitrogen metabolism in aquatic animal s is ammonia, in contrast to urea or uric acids in terrestrial animals (Wright 1995). Ammonia is present in two forms in sea water: unionized ammonia (NH3) and ammonium ion (NHt). The form of ammonia excreted by zooplankton is not certain (for crustaceans see Regnault 1987). The forms of ammonia in sea water may be expressed by the equilibrium equation: (10.2) The equilibrium reaction is influenced by temperature and pH. In the usual pH range (7.5-8.2) ofsea water and at 18 oC, less than 5% ofammonia is in the form ofNH 3. This percentage increases as temperature or pH increases. Because of the lack of information about ammonia forms excreted by marine zooplankton and complex equilibrium of ammonia in sea water, ammonia excreted by marine zooplankton has been referred as either ammonia (NH 3) or ammonium (NHt) (note that NHrN and NHt -N are comparable to each other in terms of N). In the following no distinction is made to between the two forms of ammonia. Ammonia is well documented as the major form of dissolved nitrogen excreted by marine zooplankton. For Neomysis rayii and Euphausia pacifica, ammonia-N accounted for 75%-85% ofthe total-N excreted, followed by 10%-24% as amino-N and 0%-1 % as urea-N (Jawed 1969). Ikeda and Skjoldal (1989) determined excretion rates of ammonia-N, urea-N and dissolved free amino acids (DFAA)-N of five zooplankton species (Clione limacina, Calanus glacialis, C. hyperboreus, Themisto libellula and Sagitta elegans) and noted that ammonia-N was the most important (78%-93% of the total), followed by urea-N (1 %-23%) and DFAA-N (6%-25%). Corner and Newell (1967) reported that 600/0-1000/0 of the total-N excreted by Calanus helgolandicus was ammonia and the rest was urea. Corner et al. (1976) confirmed this result for the same species fed barnacle nauplii; only 9%-10% of total-N excreted was urea. Ammonia is also the dominant form of nitrogen excreted by the ctenophore Mnemiopsis leidyi (Kremer 1977). While all these studies were made using the sealed chamber method mentioned below (Measuring metabolic rate on live zooplankton, section 10.3), Gardner and Paffenhofer (1982) determined ammonia and amino-N excretion by Eucalanus pileatus every 10 minutes with a flow-through system (see measuring metabolic rate on live zooplankton, section 10.3). According to Gardner and Paffenhofer's results, the amount of nitrogen excreted in the form of ammonia was far greater than that excreted as amino-N for this copepod. The time course of amino-N excretion was characterized by a discontinuous pattern. Although the mechanism of amino-acid release is not clear, it may come from food during feeding ('sloppy feeding'), from diffusion across cell membranes, from catabolism of food, or from osmoregulation (Gardner and Paffenhofer 1982; Forward and Fyhn 1983; Williams and Poulet 1986). In contrast to studies showing ammonia as the major nitrogenous excretory product, Webb and Johannes (1967) reported abnormally high excretion of dissolved free aminoacids by mixed zooplankton. The results ofWebb and Johannes were considered to be an artifact caused by an extremely high density of zooplankton in their experiments, i.e. amino-N leaking from the injured specimens and possible cannibalism during incubation, although Webb and Johannes (1969) suggested that the contradictory results of other workers were due to overlooking the bacterial uptake of amino-acids during the incubation (see page 471). 460 METABOLlSM Dissolved phosphorus compounds in zooplankton excreta can be separated into inorganic and organic fractions (technically, the organic fraction is not measured directly but calculated from the difference between the total and inorganic fractions). Pomeroy et al. (1963) reported that 33%-55% of the total-P excreted by mixed zooplankton was inorganic. Later, Hargrave and Geen (1968) found a high percentage in the organic fraction (up to 74% of the total-P excreted) for three copepod species. According to the seasonal study of Butler et al. (1969, 1970), the percentage of the organic fraction in the total-P excreted by Calanus finmarchicus was highest (70%) in spring when phytoplankton food was most abundant, but the inorganic fraction dominated during the food-depleted winter. It should be noted that this seasonal change in the relative amounts of organic and inorganic-P was not seen for nitrogen excretion, in which ammonia comprised a constant 88 % of the total-N excreted by the copepod. For Euphausia superba maintained with and without food for 7 months, Ikeda and Dixon (1982) found that while both organic and inorganic fractions of both nitrogen and phosphorus excreta were determined in fed specimens, only the inorganic fraction was detected for starved specimens. As was the case for nitro gen excretion, discrepancies seen in the results of phosphorus excretion experiments by previous workers may be caused by dissimilar experimental designs including the use of individual or mixed species (see page 462). Feeding conditions of zooplankton prior to or during the experiments may be considered prime factors influencing the proportion of organic fraction in total phosphorus excretion (see page 467). 1 10.3 MEASURING METABOUC RATE ON UVE ZOOPLANKTON Determination of metabolic rate is straightforward in theory and rather simple in practice. However, a number of experimental conditions are known to affect the results. Techniques currently in use for determining oxygen consumption and excretion rates of ammonia and phosphorus on live zooplankton can be classified largely into two types. 1) Sealed chamber method Specimens are confined in containers filled with sea water for a certain period and the decrease of oxygen or increase of excretory products during the period (several hours to a day) are monitored throughout, or determined at the end of the incubation. Despite the possible detrimental effects of oxygen depletion and accumulation of excreta, the method is simple in practice and has been used extensively. The technical problems associated largely with this method are reviewed in detail in the following section (Technical problems, section 10.3.1). 2) Flow-through method Oxygen consumption and accumulation of excreta are measured continuously on the specimens placed in a flow-through system. This method avoids sorne of the problems associated with method (1) but it often requires that specimens be placed in a very small cell. In addition, it requires a rigorous calculation taking into consideration flow-rate, cell size ano metabolic rates of zooplankton to obtain reliable data (Northby 1976; Niimi 1978; Propp et al. 1982). Application of this method has been limited to the study of Gerber and Gerber (1979) for simultaneous measurements ofingestion and oxygen consumption rate of tropical lagoon zooplankton, that of Gardner and Scavia (1981) for nitrogen excretion of Daphnia sp., and that of Gardner and Paffenhofer (1982) for nitrogen excretion of Eucalanus pileatus. MEASURING METABOLlC RATE ON LlVE ZOOPLANKTON 461 110.3.1 Technical problems In the following, various sources of potential error in measuring metabolic rates of zooplankton collected from the field are reviewed and discussed. For this purpose, only key references were selected. While most of the references concern zooplankton, benthic species and fishes are also referred to when appropriate. CAPTURE STRESS/STARVATION A common problem with experimental studies of zooplankton is the del ay between actual capture of specimens and subsequent measurements. Such delays may range from several hours to several tens of hours; they are often unavoidable due to the logistics of field sampling and laboratory routines. Sorne studies have noted decreases in metabolic rates of zooplankton with the increase of time after capture, and a question naturally arises as to what causes this. The validity of measurements taken at varying times after capture may be called into question as well. The first observation ofthis nature was made by Marshall et al. (1935), who reported a rapid fall in the oxygen consumption rate of Calanus finmarchicus soon after capture. Since then, similar results have confirmed this phenomenon, not only in oxygen consumption rate, but also in the excretion rates of ammonia and inorganic phosphorus of several zooplankton species (Ikeda and Skjoldal 1980; Skjoldal et al. 1984 and references therein). Compared with that of oxygen consumption rate, ammonia and phosphorus excretion rates in zooplankton just after capture are more variable due to the switch in metabolic substrates (e.g. from protein to lipid, cf. Skjoldal et al. 1984). Stress incurred during the course of sampling and possible starvation of specimens in captivity may be considered as part of the cause of the rapid fall in metabolic rates after capture (Figure 10.1). The magnitude of both effects on metabolic rates may vary from species to species, depending on the depth of occurrence of zooplankters, the sampling gear used, weight-specific metabolic rates, temperature, etc. From this scheme, a higher oxygen consumption rate just after capture may be explained by 'oxygen debt' brought about by the extra exercise during capture; this capture stress would be greatest at the beginning of an experiment. It is difficult to provide the quantity and quality of appropriate food during experiments, thereby the effect of starvation predominates during the progress of experiments. Capture stress has be en effectively demonstrated in the oxygen consumption measurements of sockeye salmon (Oncorhynchus nerka) by Brett (1964). Most of the energy needed during heavy extra exercise of sockeye salmon is supplied by anaerobic processes, not through the TCA-cycle; this is demonstrated by excessive accumulation oflactic acid in the blood (Black et al. 1962, 1966). For zooplankton, adenylate energy charge (EC) ratio has been used as an index of capture stress for the initial decreases in metabolic rate of freshly caught specimens of the copepod Acartia australis, the decapod shrimp Acetes sibogae australis (Ikeda and Skjoldal 1980), and the copepod Euchaeta norvegica (Skjoldal et al. 1984). However, no appreciable capture stress was detected from the results ofthis index in both studies. In addition, lactic acid levels in E. norvegica just after capture were low and not indicative of any oxygen debt (Skjoldal et al. 1984). Thus, the results of Ikeda and Skjoldal (1980) and Skjoldal et al. (1984) indicate that starvation was the primary cause for the decrease in metabolic rates ofthese zooplankters. A progressive decrease in metabolic rates of starved specimens for hours, days or weeks has been demonstrated in various zooplankton species (Conover 1968; Mayzaud 1976; Ikeda 1977a, 1977b; Bamstedt and Tande 1985; Kremer and Reeve 1989). The 462 METABOLlSM --- --- Time after capture Fig. 10.1 Schematic presentation of the sequential changes of the effects of capture stress and starvation on the metabolic rate of zooplankton after capture. efIect of starvation can be minimized by the use of zooplankton immediately after capture, provided there is no capture stress. Provision of appropriate food during the time between capture and experimental manipulation may be an alternative, but prolonged maintenance ofwild zooplankton in the laboratory may change its behavior, swimming activity, and nutritional condition, all of which could affect the metabolic rates. Biggs (1977) and Cetta et al. (1986) enclosed gelatinous zooplankton directly in glass jars while Scuba-diving for the measurements of oxygen consumption and ammonia excretion rates. For oxygen consumption measurements Smith (1982), Youngbluth et al. (1988) and Bailey et al. (1994) adopted a gentle suction of animal s into a chamber equipped with an oxygen sensor and incubated animals in in situ conditions on board submersibles. AH these procedures are considered to be superior to that of the conventional catch-transfer-incubation method (i.e. sealed chamber method) to minimize the efIects of handling and laboratory conditions. However, application of these methods is costly and it is difficult to control contamination by bacteria and microzooplankton in the incubation containers. Presently, in situ methods have been limited to large zooplankton which are recognizable by the unaided eye. CONTAINER SIZE/CROWDING The use of smaHer containers or high densities of specimens may be needed to bring a measurable difIerence in oxygen and/or nutrient concentrations between experimental and control chambers. Marshall and Orr (1958) examined the efIect of container size on the oxygen consumption rate of Calanus finmarchicus using various container sizes (30- MEASURING METABOUC RATE ON UVE ZOOPlANKTON 463 160 mI) and found no appreciable differences between the results. Zeiss (1963) compared oxygen consumption rates of Calanus finmarchicus and Daphnia magna placed under several densitíes of specimens. While the rates of C. finmarchicus were not affected in the range of 2.5 to 10 specimens mll, the rate of D. magna increased at higher densities. Razouls (1972) measured the oxygen consumption rate of Temora stylifera and Centropages typicus at densities from 0.1 to 2 individuals mI-l. The highest oxygen consumption rates were obtaíned at 0.5 individual mll and these rates decreased at lower and higher densities in both species. Sorne density effects have been reported in the excretion rates of nutrients by zooplankton. In Acartia tonsa the phosphorus excretion rate decreased when the density of individuals exceeded 400 specimens mll (Hargrave and Geen 1968). In contrast, Nival et al. (1974) obtained anomalously high ammonia excretion rates for Temora stylifera incubated at densities greater than 200 specimens mI-l. Container size and crowding are analogous in terms ofvolume available per specimen, i.e. density, but the former is largely interaction with container wall and the latter interaction between individuals. Therefore, the effect of these parameters on metabolic rates could vary depending on species-specific differences in swimming behavior, perception of stimuli, tolerance to a rapid depletion of oxygen, and associated accumulation of excreta. For carnivorous species, cannibalism could easily be induced by the increase in density. Thus, metabolic rates of zooplankton could be either enhanced or depressed by higher densities as described aboye. As a general guideline for designing metabolic studies, experiments using higher densities of specimens should be avoided to obtain the best results. Information about natural density of test zooplankton species should be useful to design experimental den sities of specimens for their metabolic measurements, and check the validity ofthe results whenever necessary. INJURY/DEATH Physical injury of specimens (such as broken legs or rupture of a part of the hody) incurred during collection at sea or handling in the laboratory prior to their use for experiments can easily happen, hut is often difficult to eliminate, especially for small zooplankton. Including injured or dead specimens during incubation are possible sources of error in determining metabolic rates of zooplankton. Mullin et al. (1975) found that both ammonia and phosphorus excretion of mixed zooplankton always overestimated the sum of excretion rates for healthy specimens, and considered that overestimation of the former was due to the release of nutrients by injured specimens. Ikeda et al. (1982) demonstrated that artificially injured copepods (Acrocalanus gibber and Tortanus gracilis) relea sed phosphorus at arate 6 to 9 times that of non-injured copepods (control). In the same set of experiments, artificial injury caused a twofold increase of ammonia excretion in T. gracilis, hut no significant increase was seen in A. gibber. The death of a euphausiid, Euphausia pacifica, during incubation was observed to cause an eightfold increase in phosphorus excretion when compared to healthy specimens, but little increase in ammonia excretion was observed (Table 8.3 in Omori and Ikeda 1984). Thus, it is clear that the effect of injured or dead specimens is most serious in determining phosphorous excretion rates, followed by ammonia excretion rates. The rapid release of phosphorus (PO¡-3_p) from injured or dead specimens is considered to be due to hydrolysis ofleached body ftuids. While no relevant information is available at present, reduced oxygen consumption rates are expected in experiments where injured or 470 METABOLlSM normally perform a diel vertical migration (Pleuromamma xiphias, Calanus euxinus), and found significantly higher rates in those specimens collected at night. In the same study, Pavlova (1994) noted that no day-night differences were seen in copepods in which diel vertical migration was less evident (Acartia clausi, Temora discaudata). Checkley et al. (1992) investigated diel rhythmicity in ammonia excretion rates of the migratory copepods Acartia sp. and Centropages furcatus. Specimens collected at various times within a day were used in incubation experiments (this was in contrast to the use of a stock of specimens as in the experiments of Pearcy et al. 1969; Duval and Geen 1976), and found that the daytime rates were 2.5 to 5 times greater than the night-time rates (Figure 10.3). Evaluation of endogenous diel rhythmicity in metabolic rates may not be necessary for the studies in which the major emphasis is daily metabolism (metabolic rate integrated over one day). In theory, if starvation is not a problem, this can be achieved by 12 A o 8 .i:: -b 4 &. 8o (.) ;¡¡: O + .. :J: Z o .s Q) 'ti a: 40 e B o ~ ! (.) )( w E ::J ·c ,,/ .~ o 20 E // 4 --~ E ~ --~. ~ « y .. / O 0000 0600 1200 1800 0000 Time Fig.10.3 Ammonia excretion rate of adult female copepods collected at ca. 3 h intervals for 24 h, and incubated in filtered sea water for 3-4 h under simulated in situ conditions of light and temperature. (A) Acartia sp. at 25 oc. Symbols represent duplicate ammonia analyses. (8) Centropages furcatus at 22-30 oc. Symbols: open circle, specimens from shallow warm; closed circle, those from deep warm; open triangle, those from shallow cold; closed triangle, those from deep cold waters (after Checkley et al. 1992, with permission). MEASURING METABOUC RATE ON UVE ZOOPLANKTON 471 incubating zooplankton for time periods of one half day or one full day. The diel cycles of sorne important environmental variables will then be accomplished within the laboratory experiments. The following is an example in which only cyclic changes in ambient tempera tu re are taken into account in the estimation of daily metabolismo The hyperiid amphipod Themisto japonica is a typical oceanic diel vertical migrator. During the course of its migration from the surface to 350 m depth everyday, the amphipod encounters a change in temperature from 15 oC to 1 oC in summer. Using a programmable water bath, Ikeda (1992) simulated the daily fiuctuation of temperature for migrating T. japonica, and compared oxygen consumption rates of specimens incubated in this fiuctuating temperature bath for 1 day (= 1 cycle of temperature change) with rates of those maintained in a constant temperature bath. The constant temperature bath was set to 8 oC (integrated daily mean temperature). The daily oxygen consumption of T. japonica placed in the fiuctuating temperature bath did not differ significantly from that of T. japonica in the constant temperature bath. While the associated change in hydrostatic pressure encountered by T. japonica was ignored in the experiment, previous findings (e.g. Torres and Childress 1983) suggested that effects of pressure on metabolism over the depth range of T.japonica would be minimal (see page 472). Thus, daily metabolism of migrating T. japonica can be estimated from the constant temperature experiment in which the temperature was carefully adjusted to its daily mean. Similar results were obtained for Euphausia pacifica by Torres and Childress (1983). Apparently, life in fiuctuating temperature serves mainly to eliminate the short term metabolic overshoots and undershoots typically seen in the metabolism of acclimated individual s when temperature is acutely raised or lowered (cf. Prosser 1961). BACTERIA Bacterial growth during incubation is unavoidable and may be a source of error when measuring rates of oxygen consumption andjor nutrient excretion using the sealed chamber method. Marshall and Orr (1958) used antibiotics, such as streptomycin and chloromycetin (50 mg 11 of each), to minimize bacterial oxygen consumption during their experiments on oxygen consumption and feeding in Calanusfinmarchicus. Presence or absence of antibiotics did not affect the oxygen consumption rate of the copepod, but sorne copepods cea sed feeding after addition of antibiotics. No appreciable effect of antibiotics (streptomycin, 50 mg 11) was seen in the oxygen consumption rates of Neocalanus plumchrus (lkeda 1970). For nutrient excretion experiments, Hargrave and Geen (1968) incubated several zooplankton species with and without antibiotics (penicillin and streptomycin), to measure excretion of phosphorus (inorganic and organic). In all species the excretion rates were lower for the specimens incubated without antibiotics, thus showing bacterial uptake of phosphorus during incubation. Mayzaud (1973) observed similar results for organic nitrogen excretion by Meganyctiphanes norvegica, but not in Acartia clausi in the same experimento Jawed (1969) maintained Neomysis rayii and Euphausia pacifica with antibiotics prior to nitrogen (ammonia, amino acids, urea and total nitrogen) excretion measurements in autoclaved sea water and bottles. The excretion rates of all forms of these treated specimens were not significantly different from the rates of untreated specimens. Bacteria may be either a net regenerator or consumer of nutrients, depending on their growth rate and physiological state (Goldman et al. 1987). In this light, not only the differences in zooplankton species and in experimental designs, but also the physiological condition of bacteria involved in each experiment may contribute to inconsistent 472 METABOLlSM results between studies. As in the oxygen consumption experiments aboye, the use of antibiotics requires caution concerning appropriate dosage. Sorne antibiotics interfere with the ultraviolet method for nitro gen analysis (Butler el al. 1969). Overall, the degree to which bacteria affect the measurement of nutrient excretion by marine zooplankton is presently unclear. MOLTING Molting, when it occurs during the incubation of individual crustacean zooplankton, may result in overestimates of routine metabolismo In euphausiids, molting is known to accelerate oxygen consumption rates (Paranjape 1967; Ikeda and Mitchell 1982). Bulnheim (1972) made detailed measurements of increased oxygen consumption (2.23.9 times the rate at non-molting) during the course of molting in benthic gammarid amphipods. Not only oxygen consumption, but also phosphorus excretion rate in euphausiids (Ikeda and Mitchell 1982), and ammonia excretion rate in decapod crustaceans (Regnault 1987) has been observed to increase when specimens molt during incubation. HYDROSTATIC PRESSURE Hydrostatic pressure increases progressively with increasing depth (1 atmosphere or 101.3 kPa per 10 m depth). Early interest in the effect of hydrostatic pressure on zooplankton metabolism mainly concerned di el vertical migrators, particularly residents ofthe deep scattering layer that experience large pressure changes (~20 atm) twice daily as a result of their vertical excursions. Oxygen consumption rates of di el vertical migrators such as euphausiids (Te al and Carey 1967; Pearcy and SmallI968), decapods (Teal 1971), and thecosomatous pteropods (Smith and Teal 1973) determined under various combinations of temperature and hydrostatic pressures in the laboratory yielded a consistent result that the rates were affected by temperature but not by pressure. The lack of appreciable hydrostatic pressure effects on oxygen consumption rates has recently been confirmed on bathypelagic chaetognaths, hydromedusae and a polychaete by comparing rates at 1 atm and at 100 atm in the laboratory (Childress and Thuesen 1993). All these results suggeSt that estimation of oxygen consumption rates of zooplankton living at depth is experimentally feasible at 1 atm, provided that livespecimens are recovered successfully to the surface without damage. In this regard, Bailey el al. (1994) noted a 10ss in motor activity of delicate mesopelagic gelatinous zooplankton due to decompression, which was reflected in reduced oxygen consumption rates. It is well documented that the metabolism of pelagic crustaceans, fishes and cephalopods declines with the increase of depth of occurrence (Childress 1975; Quetin el al. 1980; Ikeda 1988; Torres et al. 1979, 1994; Childress 1995). This depth-related decline of crustacean and fish metabolism is not due to the increase of hydrostatic pressure as mentioned aboye, nor to the decrease of temperature; it is an adapted characteristic ofmany species of deeper-living fauna (Childress et al. 1980; Childress and Micke11985; Ikeda 1988). A comparison of oxygen consumption rates and intermediary metabolic enzyme activities in chaetognaths living at various depths by Thuesen and Childress (1993a, 1994), revealed that a depth-related decline in metabolism was not the case for chaetognaths and medusae. Thuesen and Childress (1993a, 1994) and Childress (1995) postulate that a depth-related change of metabolism would be selected for in visual predators that require a well-developed musculature to chase down prey that can be visually targeted in well-lit surface waters, but not in visual predators in the light- MEASURING METABOUC RATE ON UVE ZOOPLANKTON 473 limited deep-sea. The limited distances for visual detection afforded by the low light levels at depths inhabited by deeper living species has resulted in a loss of locomotory musculature and a concomitant decline in metabolic rates. The same argument does not apply to non-visual predators such as chaetognaths whose methods of prey detection are not appreciably altered with depth. In light of the diversity of pelagic fauna in the deepsea, more data are needed to prove or disprove the present hypothesis for reduced metabolic rates of deep-living zooplankton. TURBULENCE In the sealed chamber method for measuring metabolic rate, zooplankton are typically confined in standing (turbulence-free) water during the experiment. In nature this is not the case, and zooplankton living in the shallow layers of the sea are more or less under the infiuence of small-scale turbulence at all times. Small-scale turbulence can affect zooplankton metabolism in two ways: (1) by increasing the encounter probability between zooplankton and food particles, and (2) by increasing the frequency of escape reaction of zooplankton, a metabolically expensive swimming behavior (Saiz and A1caraz 1992). Saiz and Alcaraz (1992) placed Acartia sp. in 25to 50-mI screw-cap polyethylene vials three-quarters filled with GF /F filtered, air-saturated sea water, and the vials were placed on a reciprocal shaker (90-100 strokes minl, 2.5 cm amplitude) for 24 h to determine their ammonia and phosphorus excretion rates. As a control, Acartia sp. was placed in standing vials. The excretion rates of Acartia sp. placed on the shaker were 1.6 times higher on average than the rates of controls. Saiz and Alcaraz (1992) noted that the intensity of turbulence generated by shaking in their experiments was not quantified, but was probably higher than that found in the fie1d, at least in oceanic waters. Clear1y, oxygen consumption is superior to nutrient excretion for evaluating the effects of smallscale turbulence on zooplankton metabolismo In a later study, A1caraz et al. (1994) determined heartbeat rates, instead of oxygen consumption rates, of several zooplankton species (both marine and freshwater) under quantified turbulence (ca. 5 mm2 S-3) and observed increases ranging from 5% to 93%. However, a poor correlation between heartbeat and oxygen consumption rates was reported for Euphausia superba and Parathemisto gaudichaudi by Opalinski (1979). More investigation is needed to evaluate and generalize the effect of small-scale turbulence on zooplankton metabolismo Clear1y, development of new experimental techniques and quantitative evaluation of turbulence levels encountered by zooplankton in the field will help to resolve the interactions between turbulence and metabolism in zooplankton. 110.3.2 Body size and temperature as bases of metabolic comparison Metabolic rate (M) of animals is known to vary as a function of body mass (W) intraspecifically and interspecifically (Zeuthen 1947, 1953; Hemmingsen 1960). The relationship is expressed as (10.5) where a and b are constants. According to Zeuthen's (1953) recapitulation theory the mas s exponent b varies successively with the increase ofbody mass, from 0.75 to 1.0, then to 0.75, where the middle 1.0 is typical in very small metazoans phylogenetically or middevelopment stages ontogenetically. However, Banse (1982) re-analyzed published data ofmetabolism-body mass relationships ofvery small invertebrates, concluding that the b 474 METABOLlSM for this group ofanimals was nearO.75 instead of 1.0. Thus, Zeuthen's theory appears not to be warranted any more (Banse 1982). For marine planktonic metazoans ofwhich body mass ranges five to six orders of magnitude at most, the mass exponent b is characterized as less than 1 and usually in the range ofO.7 to 0.9. Within narrow body mass ranges the relationship between metabolic rate and body mass is often masked by the scatter of the data, i.e. the confidence interval of b is too wide to judge b < 1. From a statistical viewpoint, a geometric (GM) regression model, rather than arithmetric (AM) regression model (= the least-square regression), is appropriate for calcu1ating b (Ricker 1973; Laws and Archie 1981). A body mass exponent calculated from AM regression (b) and that (d, i.e. M = e Wd, where e and d are newly designated constants) from GM regression are the same when the correlation coefficient (r) is 1.00, but the former is greater than the latter when r < 1.00, and r < 1.00 is usually the case (i.e. d = b/r cf. Ricker 1973). For metabolic comparison between species with dissimilar body masses, a common constant b needs to be established or assumed (or else the conclusion will vary depending on the choice ofbody mass). The results ofmetabolic comparison are also affected by the expression of body mass when the body composition of the species to be compared is different. Ikeda and Mitchell (1982) compared oxygen consumption rates of Euphausia superba and Salpa thompsoni on the basis of equivalent body mass and noted that the rate of the former was greater than the latter by a factor of 17 on a wet weight, 4 on a dry weight, Snd 0.8 on a carbon or nitrogen basis. Carbon or nitro gen units appear to be superior to wet and dry weight to reduce the phylogenetic differences in body composition for the purpose of interspecific metabolic comparison. Schneider (1990) reached the same conclusion from the body carbon based comparison of ammonia excretion rates between gelatinous and nongelatinous zooplankton. Ikeda (1988) proposed 'Adjusted Metabolic Rate' (AMR) for metabolic comparison between mesopelagic and epipelagic zooplankton. AMR is defined as M divided by (body nitrogen)-b. The mass exponent b used in the comparison oflkeda (1988) was 0.8505 for oxygen consumption rate, 0.8361 for ammonia excretion rate, and 0.8704 for phosphorus excretion rate, all of which are derived from statistical analysis of comprehensive data sets describing epipelagic zooplankton metabolism collected from several regions of the world ocean (cf. Table 10.2). The relationship between metabolic rates (M) and temperature (TOC) is frequently described by QlO (se e Temperature, section 10.3.1). When the QlO is constant over a given temperature range, the relationship is re-expressed as M = apT, where a and pare constants. Since it is known already that temperature effects are difficult to analyze using laboratory acclimation experiments (see Temperature, section 10.3.1), a comparison of metabolic rates of zooplankton living in dissimilar thermal regimes is most appropriate for drawing generalized conclusions regarding temperature effects. Intraspecific comparisons would be ideal, but the data presently available are not sufficient for this analysis. Interspecific comparison is the only alternatíve. Ikeda (1985) compiled oxygen consumptíon data of 143 zooplankton specíes, ammonía excretion data on 131 species, and phosphorus excretion data on 52 species (general size range: 103 to 10 3 mg dry weíght) from tropical, sub tropical, temperate, subarctic and Antarctic waters (temperature range: -1.4 to 30 OC) and analyzed the data as a function ofbody mas s and habitat temperature, assuming a constant mass exponent b in metabolism-body mass relationship and a constant QlO over the temperature ranges investigated. His results revealed that the 940/0 to 95% of the variation in oxygen consumption rates could be ascribed to habitat temperatures and body sizes of zooplankters (Table 10.2). Excretion rates of ammonia and inorganic phosphorus were Table 10.2 Regression statistics of metabolic rates (Y= oxygen consumption, ammonia excretion, or inorganic phosphorus excretion rates) on body mass (X 1 = dry (DW), carbon (CW), nitrogen (NW), or phosphorus (PW) weight) and habitat temperature (X 2) for marine zooplankton. 010 calculated from a2 are shown on the right-hand side of the table (010 = exp(1 O x a2» (modified from Ikeda 1985). " In Y=ao + I 81 In Xi + B2X2 Metabolic rate Body mass unit N ao a1 ~ ,¡z Q10 Oxygen uptake DW 721 -0.2512 0.7886 0.0490 93.9 1.632 CW 721 0.5254 0.8354 0.0601 95.5 1.824 NW 721 1.7412 0.8505 0.0636 95.1 1.889 PW 721 3.7890 0.8167 0.0552 94.0 1.737 Ammonia excretion DW 1186 -2.8900 0.7616 0.0511 85.4 1.667 CW 1186 -2.1763 0.8293 0.0648 86.5 1.912 NW 1186 -0.9657 0.8361 0.0656 86.2 1.927 PW 1186 1.0708 0.8063 0.0562 84.5 1.754 Phosphate excretion DW 749 -4.3489 0.7983 0.0285 86.4 1.330 CW 749 -3.6031 0.8622 0.0438 86.5 1.550 NW 749 -2.3490 0.8704 0.0441 86.9 1.554 PW 749 -0.1839 0.8569 0.0376 84.2 1.443 476 METABOLlSM more variable; 84% to 87% of their variability was explained by changing temperature and body mass. Because of the wide variability in the characteristics of the studied zooplankton species, QlO varied as a function of the body mass unit chosen for comparison. General ranges were 1.63 to 1.89 for oxygen consumption rate, 1.67 to 1.93 for ammonia excretion rate, and 1.33 to 1.55 for phosphorus excretion rateo The combination of body mass-specific Q10 with AMR can allow standardization of metabolic rates in terms of body mass and temperature as bases for metabolic comparison of zooplankton. The data analyzed in Table 10.2 do not inc1ude metabolic data of Arctic zooplankton. Because of the interest in historical 'metabolic cold adaptation', oxygen consumption rates of Arctic species are of special interest, i.e. despite a similar sub-zero habitat temperature the rates of Arctic zooplankters are expected to be lower than the rates of Antarctic zooplankters because the Arctic ecosystem is younger. For a detailed historical account of this hypothesis see Holeton (1974) and Clarke (1983). An intra-generic comparison of oxygen consumption rates in selected zooplankton species from the Arctic and Antarctic revealed no significant difference between the two (Ikeda 1989a). Thus, the regression equation in Table 10.2 can be used as the basis for metabolic comparison of zooplankton living over world oceans. As an obvious divergence from the regression equation in Table 10.2, extremely reduced metabolic rates have been reported for overwintering copepods in 'diapause'; i.e. Calanus pacificus californicus in the Santa Barbara Basin (Alldrege et al. 1984), Calanusfimarchicus and C. helgolandicus in Norwegian fjords (Hirche 1983), Calanoides acutus in the Weddel Sea (Drits et al. 1994), Neocalanus cristatus, N. plumchrus and Pseudocalanus minutus in the Japan Sea (Ikeda and Hirakawa 1998). These copepods are primarily herbivores living in cold water, and their diapause stages (mostly late copepodite stages) are characterized by their occurrence from the mesopelagic zone, motionless in behavior and a large accumulation of lipids (stored energy) in the body. For a recent account of dormancy (inc1uding diapause) of planktonic copepods, see the review ofWilliams-Howze (1997). 110.3.3 Metabolic quotients Simultaneous measurements of oxygen consumption, ammonia excretion and inorganic phosphorus excretion allow the ratios of oxygen consumption to ammonia excretion (O:N), ammonia excretion to inorganic phosphorus excretion (N:P), and oxygen consumption to inorganic phosphorus excretion (O:P) to be compared on an atomic basis. These ratios, called metabolic quotients, change depending on the metabolic substrates of the animal. Nitrogen and phosphorus contents of animal protein, lipid, and carbohydrate, together with the amount of oxygen required for combustion of each c1ass of biomolecule are summarized in Table 10.3. The O:N, N:P and O:P ratios for each c1ass of organic matter are also ca1culated. It must be noted that these ratios will change slightly, because of diversity within a c1ass of biomolecule, particularly in the make-up of protein and lipid, and no precise information is available for zooplankton material. Even so, consistent variations are observed in these ratios depending on whether the metabolic substrate is protein, carbohydrate or lipid. It is apparent from Table 10.3 that a carbohydrate-dominated metabolism causes high O:N and O:P ratios. Low N:P and O:N ratios are characteristic oflipidand protein-oriented metabolisms respectively. Raymont and Krishnaswamy (1960) and Raymont and Conover (1961) observed little MEASURING METABOUC RATE ON UVE ZOOPLANKTON 477 Table 10.3 Average nitrogen and phosphorus composition of organic matter, the oxygen required to oxidizeeach class of organic matter in an animal body, and calculated metabolic quotients (O:N, N:P and O:P, by atoms) (modified from Ikeda 1974). Carbohydrate lipid Protein N (g/g)a O 0.0061 0.178 P (g/g)a O 0.0213 0.007 O (I/g)b 0.83 2.01 1.02 O:N 00 412 7.2 N:P 0.63 56 O:P 00 261 403 a From Rogers (1927) b From Gnaiger (1983) change in the carbohydrate content of starved zooplankton. In conjunction with these observations, the carbohydrate content of zooplankton was found to be only a few percent ofthe dry weight (5% at most). Even if all the carbohydrate in a zooplankter was metabolized, it would not be large enough to support the animal's metabolic requirement for one day. Therefore, protein and lipid are considered the major metabolic substrates of starved zooplankton. According to Conover and Corner (1968), the O:N ratio of sorne boreal zooplankton is high at the end of summer but declines throughout winter, with a corresponding decrease in lipid. During the spring phytoplankton bloom, zooplankton feed actively on phytoplankton and store lipid, but the O:N ratio remains low during this periodo The O:N ratio increases only after the zooplankton have deposited a large amount of lipid, indicating a close association of the O:N ratio to the lipid content of zooplankton. Assuming that the respective nitro gen content of protein and lipid is 16% and 0%, and the oxygen required for complete combustion per 1 g for protein and lipid is 1.04 and 2.02 1, Ikeda (1974) calculated an O:N ratio of 24 when protein and lipid are metabolized in equal quantities at the same time; hence an O:N ratio less than 24 indica tes protein-oriented metabolism and a ratio greater than 24 indicates lipidoriented metabolism (from the data in Table 10.3 this is re-calculated to be 20.6). O:N ratio s were used to compare the metabolic substrates of zooplankton inhabiting a large geographical area extending from tropical through subarctic waters. As a result, he concluded that zooplankton living in tropical, subtropical and temperate waters were characterized by protein metabolism, while those living in subarctic waters showed a wide range in the O:N ratio. In addition to season (Conover and Corner 1968) and specific differences (lkeda 1974; Ikeda and Mitchell 1982; Gaudy and Boucher 1983; Quetin et al. 1980; Ikeda and SkjoldalI989), O:N ratios are known to change as a result ofstarvation (Mayzaud 1976; Ikeda 1977b), recent feeding conditions ofanimals (lkeda and Dixon 1984), and acclimated temperature (Mayzaud and Conover 1988). Extremely low O:N ratio s (lower than the theoretical minimum of 7.2, see Table 10.3) seen in the results of sorne workers in Table 10.4 may be caused by artificially high excretion rates of animals, the result of severe stress either in the field or in the laboratory (Mayzaud and Conover 1988). Mayzaud and Conover (1988) reviewed the O:N ratio in marine zooplankton and noted that while the O:N ratio reflected changes in the biochemical composition of the body for zooplankton with large energy reserves (lipid), the ratio for Table 10.4 The O:N, N:P and O:P atomic ratios from the measurements of oxygen consumption, ammonia excretion and phosphorus excretion rates being reported by previous workers.a Zooplankton O:N N:P O:P References Mixed zooplankton (UFW) 7.7 7.0 54.0 Harris (1959) Mixed zooplankton (UFW) 41.0 9.98 222.0 Martin (1968) Calanus cristatus (UFW) 5.7 19.0 110.0 Taguchi and Ishii (1972) Calanus plum eh rus (UFW) 6.8 13.0 89.0 Taguchi and Ishii (1972) Mixed zooplankton (UFW) 13.48 10.33 142.4 Le Borgne (1973) Sagitta hispida (FW) 11.3 Beers (1964) Mixed zooplankton (UFW) 72.0 Satomi and Pomeroy (1965) Calanus helgolandicus (FW) 9.8-15.6b Comer et al. (1965) Boreal zooplankton (10 species) (UFW) 6-200b Conover and Comer (1968) Calanus finmarehieus (FW) 10.8 Butler et al. (1969) Calanus finmarehieus (FW) 11.0c Butler et al. (1970) Sagitta hispida (FW) 6.8 Reeve (1970) Calanus helgolandieus (FW) 16.5c Comer et al. (1972) Temora stylifera (FW) 7-15d Nival et al. (1974) Mediterranean zooplankton (4 species) (FW) 1.6-12.1 Mayzaud (1973) Boreal, temperate, subtropical and tropical 4-115 Ikeda (1974) zooplankton (81 species) (FW) Mixed zooplankton (UFW) 6.8 Mullin et al. (1975) Mesopelagic crustaceans (14 species) (FW) 9.1-91.0 Quetin etal. (1980) Antarctic zooplankton (14 species) (FW) 7.0-19.8 2.5-24.7 43-304 Ikeda and Mitchell (1982) Tropical zooplankton (27 species) (FW) 2.7-28.6 2.3-44.5 19.6-410 Gaudy and Boucher (1983) Calanus glacialis (FW) 3.0-9.9 7.7-12.9 38.7-76.5 Bamstedt and Tande (1985) Tropical ctenophores (4 species) (FW) 10.2-15.8 Kremer et al. (1986) Salps (10 species) (UFW, FW) 13-28 Cetta et al. (1986) Antarctic mesopelagic zooplankton (7 species) (FW) 14.1-73.9 1.8-15.9 67-290 Ikeda (1988) Arctic zooplankton (10 species) (FW) 7.0-19.8 2.5-24.7 43-304 Ikeda and Skjoldal (1989) a Use of unfiltered (UFW) and filtered (FW) sea water for measurement is noted. -= no data. b Ninhydrin N. e Total N/total P. dTaken from Nival et al. 1974, Figure 10. METABOLlC RATE ANO ENZYMATIC INOICES 479 small neritic zooplankton with few energy reserves reflected the qua lit y and quantity of food in the environment. The N:P and O:P ratio s are not as sensitive as the O:N ratio for assessing metabolic substrates of zooplankton, though in earlier studies interest centered around the constancy of N:P and O:P values. Satomi and Pomeroy (1965) compared the O:P ratio ofmixed zooplankton from three different sites and obtained quite similar values (63-75, with a mean of 72). From a seasonal study of total nitrogen and total phosphorus excretion by Calanusfinmarchicus, Butler el al. (1970) found the atomic ratio ofN:P was stable through spring (11.0) and winter (14.6). If the N:P and O:P ratios are constant, estimation of oxygen consumption, nitrogen excretion and phosphorus excretion rates can be made by measuring only one of these variables. However, subsequent studies by other workers revealed that there is a large variation in these ratio s (Table lOA). The N:P and O:P ratios are now known to change not only between species (Ikeda and Mitchell 1982; Gaudy and Boucher 1983; Ikeda and Skjoldal 1989) but also within the same species, depending on temperature (Le Borne 1982), time for incubation (Le Borne 1979; Ikeda and Skjoldal1980; Bamstedt and Tande 1985), quality and quantity offood given (lkeda and Dixon 1984), and prolonged starvation of animals over weeks (Ikeda 1977b). Since an abnormally high phosphorus release has been observed in damaged zooplankton (see Injuryjdeath in section 10.3.1), extremely low N:P and O:P ratio s may be used as indicators of damage in experimental animals. Redfield el al. (1963) proposed the average C:N:P composition ofmarine phytoplankton and zooplankton as 106: 16: 1 by atoms. Richards (1965) constructed a model of the organic matter composition of phytoplankton and zooplankton based on Redfield's ratio and a sequence of decomposition: (CH20)106(NH3)16H3P04 + 1060 2 = 106C0 2 + 16NH 3 + H3 P0 4 + 106H 20 (10.6) 16NH 3 + 320 2 = 16HN0 3 + 16H 2 0. (10.7) Hence, (CH20)106(NH3)16H3P04 + 1380 2 = 106C0 2 + 122H 20 + 16HN0 3 + H3 P0 4. (10.8) From this scheme, the O:N, N:P, and O:P ratio s are predicted to be 17, 16, and 276, respectively. When biological oxidation by zooplankton is considered, the nitrogenous end product is not NH0 3 but NH 3. Thus, the appropriate ratios would be 13, 16, and 212. Application of these ratios to individual zooplankton species is cautioned, since a large departure ofthe C:N:P ratio from that ofRedfield has been noted (cf. Corner and Davies 1971). 1 10.4 METABOLlC RATE ANO ENZYMATIC INOICES The search for physiological indices to characterize field-caught specimens has been slowly gaining momentum in biological oceanography for the last 30 years. Physiological indices may be thought of as biochemical proxies for a physiological rate that is either difficult to measure or where time constraints make sufficient data collection difficult in the time frame of a single cruise. The development of reliable enzymatic indices will help resolve the large hysteresis between the data gathering efforts of physical and chemical Q) -- cu..c ~-.... co..c :.¡:o O) 0..'- E ~ ::l_ en Q) § ~ (0) c-.... Q) C\I 0)0 ~ X::J.. 0A 0.097 ~ 0.087 .. 0.077 ~ 0.067 - 0.057 - 0.047 ro 0.037 I I 1.24 r I -- . -- 2.06 3 ZONE I 0.65 -- 5 . - - - - ..c O) Q) ~ - Q) ~ B 2.4 ;. . 2.0 .. 1 -- . 1.6 0.65 1.2 - -- - - 0.8 ro 2.06 1 1.24 O~----~I------.'------l-------- 1 3 5 ZONE Fig.10.6 Oxygen consumption rate and citrate synthase (es) activity, the enzyme catalyzing the initial step in the Krebs eycle, of the Antarctic copepod, Rhincalanus gigas, with relation to the Antarctic ice edge. Zone 5 is consolidated pack-ice, zone 3 is the highly productive marginal ice zone, zone 1 is the open water seaward of the ice-edge bloom. Oxygen consumption rates are expressed as JlI O2 9 wet weight1 h -1. Enzyme activity is expressed in international units per gram wet weight (lU 9 -1: IU = Jlmol substrate converted to product per minute). Error bars are 95% confidence limits. Numbers within the figure are the values for chlorophyl biomass (Jlg 1-1) corresponding to each zone. Oxygen consumption rates were significantly (P < 0.05; ANOVA) lower in the pack ice (zone 5) than at the ice-edge (zone 3) or open water (zone 1) locations. es activities were significantly (P < 0.05; ANOVA) lower in the pack ice (zone 5) than at the ice-edge (zone 3) or open water (zone 1) locations. Oxygen consumption data contributed by Helena Kawall, and es activity data by Steve Geiger. METABOLlC RATE ANO ENZYMATIC INOICES 487 (Kenner and Ahmed 1975). As mentioned in Enzymes of intermediary metabolism, section 10.4.2, Clarke et al. (1992) found that growth of ichthyoplankton (larvae of Sciaenops ocellatus and Lutjanus synagris) correlated well with activities of LDH and CS. In contrast, Berges et al. (1993) observed that activities of CS and GDH did not corre1ate well with specific growth rate in laboratory-raised Artemia franciscana and cautioned against their use in the field. These inconsistent results may refiect the physiological dissimilarities of experimental organisms between studies, but evaluation of growth conditions may be a necessary adjunct for obtaining reliable correlations between metabolism and biochemical proxies in field-caught plankton. BOOY SIZE A series of two papers (Berges et al. 1990; Berges and Ballantyne 1991) described scaling of metabolic enzyme activities with body size at different ration levels and in different species (Artemiafranciscana, Daphnia magna). The authors concluded that scaling with body size was enzyme dependent and often differed from the classic 'b-value' of 0.75 in the allometric equation between metabolic rate (M) and body mass (W) (i.e. M = aW b, where a is a constant, see also Body size and temperature as bases of metabolic comparison, section 10.3.4, and equation 10.5), hence scaling must be considered in any field application of enzyme activities as biochemical proxies. A large amount of scatter in the R/ETS and GDH/NH 4 data in Figure 10.4 may be due to the mixture of zooplankton with different body sizes (= body mass). Christensen et al. (1980) compared R/ETS ratios of various marine organisms including bacteria to zooplankton (body mass: 107 to 10 3 mg dry weight) and noted only a slight dependence of the ratio on the body mass (mg DW) of organisms: IOglO(R¡ETS) = -0.0485Iog 10 DW + 0.191. (10.9) Packard (1985) summarized the R/ETS ratio to be 0.17 for phytoplankton, 1.1 for bacteria, 0.25 for protista and 0.49 for zooplankton. Analyzing the relationship between respiration rates and ETS activities, a linear regression, instead of a mean, has been suggested to improve the precision in estimation of the former from the latter (King and Packard 1975; Arístegui and Montero 1995). MISMATCH BETWEEN METABOLlC RATE ANO ENZYME ACTIVITY Bamstedt (1980) noted a delayed response ofETS activities to increased respiration rates in Acartia tonsa. Finlay et al. (1983) found a time-lag between respiration rates and ETS activities on ciliates which were starved and then fed. Finlay et al. explained the cause of the time-lag as a limitation in the amount of enzymes involved in the ETS. This uncoupling would be a source of variation in the relationships between enzymatic activities and metabo1ic rates of organisms collected in the field, where food availability may change over short time scales as a consequence of physical instabilities of water columns. The time-lag observed by Finlay et al. (1983) on ciliates was on an hourly time scale. While the application is presently limited to the bacteria bisubstrate enzyme kinetics model proposed by Packard et al. (1996) should provide a better insight to the uncoupling problems between respiration and ETS activity. CALlBRATION OESIGN Present calibration procedures for the relationship between enzyme activity and metabolic rate appear inadequate. Figure 10.7 illustrates a typical calibration of ETS activities against respiration rates using the sealed chamber method as practiced by 25~--------------------------------~ 20 ... - ..c ca E 15 e ca - 0(\1 10 ::l.. 5 A ETS ~- RI ,--- .. ............. ~---- " ..... ""'- - ........... ...... - ...... R O~----------------------------------~ 1.2 1.1 1.0 0.9 0.8 0.7 0.6 0.5 B RI/ETS R/ETS ----------------- 0.4 4__ --~__._--_r___r_--.,....____r_--_r_____r_--r__--r----t O 5 Incubation time (h) 10 Fig.10.7 (A) Model run for the likely changes in the ETS activity and respiration rate (R, and its integrated mean over incubation time R') of zooplankton placed in a closed bottle filled with filtered sea water. (B) Resultant R/ETS (and R' IETS) ratio. In this run, ETS and R at the start (t= O) is designated as 20 and 10,u1 O2 (animal h)-" respectively, and decrease exponentially to 4 and 2,u1 O2 (animal h)-1, respectively, at the end of incubation lasting for 12 h, maintaining a constant R/ETS ratio of 0.5. Note that the decrease with the progress of incubation time of R' is much'slower than that of R, which reflects an increase of R' IETS during the incubation. Assumed equations for the change with time are; ETS = 20;O.1341t, R = 10;O.1341t, and R'= J¿ Rdt= 10(1 - e-O. 1341t )/(0.1341t). CONCLUDING REMARKS 489 previous workers (Bamstedt 1980; Ikeda and Skjoldal1980; Skjoldal el al. 1984). In the sealed chamber method, respiration rate is determined from the difference in the oxygen concentration between experimental and control bottles at the end of the incubation (i.e. R': integrated mean respiration rate over the incubation period in Figure 10.7), and ETS activity is measured on zooplankton retrieved from experimental bottles at the end of incubation. Because of this methodological constraint, the R'jETS ratio obtained from the sealed chamber method is subjected to error unless there is a constant respiration rate and ETS activity over the incubation periodo In the model shown in Figure 10.7, a rapid decrease of ETS activity and a less pronounced decrease of integrated mean R' reflected a progressive increase of the R' jETS (1.24 at the end of 12 h, in contrast to a designated ratio of 0.5). The problem highlighted in the model is not limited to the ETS activity, it applies to the activity of other enzyme systems as well. To overcome this problem, a continuous recording system for metabolic rate, such as the flow-through system (Measuring metabolic rate on live zooplankton, section 10.3) is more appropriate for the purpose of calibration. 1 10.5 CONCLUDING REMARKS While the determination of metabolic rates of live zooplankton is easily accomplished with either the sealed chamber method or the flow-through method, extrapolation of measured rates to a species population in the field is not straightforward. One obvious problem is the lack of a cIear index to judge whether or not the specimens collected from the field and brought into the laboratory are 'damaged'. Part of the problem may be overcome by experience and by checking morphology, coloration, swimming behavior, orientation patterns, and response to stimuli (light, touching with needle, etc.). However, this sort of quick screening prior to an experiment may not always detect physiological damage. The second problem associated with experiments on live zooplankton is the difficulty in reproducing natural environmental conditions for zooplankton in a laboratory experimento Environmental and nutritional conditions of zooplankton in the laboratory affect the metabolism of zooplankton. Our knowledge about the quantitative metabolic responses of zooplankton to environmental and nutritional conditions is quite limited. Most previous studies concern one or two potential parameters affecting metabolism; few attempts have been made to examine the combined effects of various parameters. Newell el al. (1976) analyzed a combination of six variables (exposure temperature, body weight, starvation period, feeding period, acclimation temperature, and lipid content) on the oxygen consumption rate of an intertidal isopod, Ligia oceanica, in the laboratory. They found that 85% ofthe observed variation in the rates could be attributed to the six parameters. An approach of this sort is of great importance for the extrapolation of laboratory results to field populations of zooplankton. The third problem is the lack of adequate techniques to control swimming activity of test zooplankton during experiments. It has been assumed that the change in swimming activity of animals brought in the laboratory and confined in containers for metabolic measurements is minima!. This may be the case for small, less active zooplankton. For larger and more active zooplankton such as euphausiids, determination of metabolic rate under well defined activity levels of animal s (i.e. standard, routine, active metabolism) is needed for valid comparison of the data between workers and for better estimation ofmetabolic costs for diel vertical migration in the field (Torres and Childress 1983). 490 METABOLlSM The fourth problem is associated with zooplankton themselves; an assemblage composed of highly diverse groups of animals (11 phyla at least, cf. Omori and Ikeda 1984) poses a real problem in establishing standardized methods for measuring their metabolic rates. High phylogenetic diversities are inevitably accompanied by a wide body-size range, and dissimilarities in behavior, locomotory activity, physiology and nutrition of species are involved. It is almost certain that inconsistent experimental results on zooplankton between workers reflect the complexities of the animal groups they have studied. Contrary to earlier expectations, enzymatic methods for estimating metabolism are not very simple, and are not necessarily free from the problems discussed here. In addition to requirements for proper calibration with metabolic rate measurements on live zooplankton under controlled laboratory conditions, information about nutritional state, growth condition and size of animals may also be required for the better estimation ofmetabolic rates ofzooplankton in the field. In this regard, direct determination (sealed chamber method or flow-through methods) and indirect determination (enzymatic methods) will complement each other as we work toward establishing better methodologies for assessing zooplankton metabolismo Practice (T. Ikeda and J.J. Torres) 1 10.6 COLLECTION AND HANDLING OF ZOOPLANKTON Most zooplankters are fragile, and for use in metabolic experiments it is important that the specimens remain undamaged. Figure 10.8 shows two examples of collecting buckets designed for such samples (see also Reeve 1981), their volumes (2 to 30 1) are several times larger than that of an ordinary cod-end bucket and they have mesh windows in the anterior part. The cod-end bucket is filled with sea water prior to sampling so that the net will sink promptly when deployed. A vertical haul or short horizontal tow with a fine-mesh net is preferable in order to avoid damage to the plankton during the haul. The towing speed NET NET NET CANVAS NET .¡ I .. :~ fi ·····U;ll I I I I POL YETHYLENE BOTTLE '., ) WI11-OOT BOTIa-1' • Fig. 10.8 Two examples of eod-ends for eolleeting live zooplankton (after Omori and Ikeda 1984, 'Methods in marine zooplankton eeology' (ISBN: 047/-80/070), John Wiley & Sons, Ine., reprinted by permission of John Wiley & Sons, Inc.). COLLECTION AND HANDLING OF ZOOPLANKTON 491 should be about 0.3 m s -1 or less, and when sampling from a large ship, it is better to stop the engine and make use of the drift of the ship caused by the wind. A long tow weakens the plankton in the neto In tropical waters, where the water temperature is high, very short tows should be used. After retrieving the net, the bucket is quickly placed in a container filled with a large amount of sea water, after which the samples are gently transferred to the container. The plankton that remain attached to the net should be considered damaged and not transferred into the container. As plankton are damaged from direct exposure to the sun, the best sampling time is after sunset (Light, section 10.3.1). To sample actively swimming macroplankton, a net with a large mouth and good filtration efficiency, such as the ORI-C net (Omori 1965), should be used. The posterior part of the ORI-C net is made of mesh gauze which is finer than the anterior section to minimize damage to the organisms during the tow. Additional protection for specimens may be achieved by using specialized cod-end buckets that are designed to c10se at the end of the tow, thereby creating a parcel of protected water for bringing animals to the surface unharmed. For example, the thermal cod-end described by Childress et al. (1978) has been used to successfully recover deep-living oceanic zooplankton. Johnson and Attramadal (1982) devised another type ofprotective cod-end bucket for use on sledges to collect epibenthic animals. This simply designed bucket can be modified for midwater horizontal tows to avoid the mechanical damage of specimens and prevent exposure to large fluctuations in temperature, salinity, and light during recovery. Gelatinous plankton are best captured by divers who introduce them into wide-mouth containers, but sorne may be obtained with net hauls in the manner described aboye. To transfer specimens to another container, they should be scooped gently with a small beaker or specially designed net (Figure 10.9A). When transporting the living zooplankton to the laboratory, their density should be kept as low as possible. Polyethylene containers with airtight lids are excellent for this purpose. To reduce damage of organisms due to agitation of water during transport, the container should be completely filled. AIso, every effort should be made to avoid direct sunlight and change in water temperature. It is therefore advisable to cover the containers with dark c1oth. For certain species aeration (gentle bubbling of sea water with air from air compressors) may be needed iftransportation takes a long time. For sorting specimens, a portion of the sample is gently poured into a transparent container of appropriate size and healthy animals are removed by pipette, spoon or scoop. The pipettes are made of glass or c1ear acrylic tubes of various diameters. Simple apparatus and methods to aid sorting of live zooplankton are shown in Figure 10.9 (B and C). In sorne cases the sample may be left for 10 to 30 min after having been transferred to a container, and the differences in the phototaxis or geotaxis of different species used to good advantage in sorting. Small transparent organisms are easily seen if the background of the container is darkened and light enters through the side. For sorne species, their characteristic swimming behavior as well as body coloration may distinguish them for sorting. Selection of experimental specimens is based on appearance. Any evidence of physical damage such as bruising or opaque patches on an otherwise transparent individual constitutes grounds for rejection. With crustaceans, any fouling of limbs or unnaturally limited range of motion on the swimming legs would be a basis of rejection. It is worth the energy to observe living zooplankton in beakers, buckets, or petri-dishes to get a feeling for how they move and what a normal range of limb motion is. There is no substitute for knowing your animal! 492 METABOLlSM A B y Netting Plastic bag r e Sample 1_ _ _ _ _ _ Lower filtering tube - ... t!ii I .~ Fig.10.9 Simple apparatus to aid sorting of living zooplankton. (A) net of which the lower part is replaced with a plastic bago With this net large zooplankton are transferred with a certain amount of water from one container to the next without exposing them to air. (B) A siphon system to remove unwanted small specimens. On one end of the system is fitted a funnel covered with a mesh screen. A gentle siphoning out of the water is essential to avoid damaging larger specimens. (C) A double filtering system to remove undesired larger and smaller zooplankton simultaneously. A zooplankton sample is gently poured into the upper filtering tube through a coarse mesh screen and passes into the lower filtering tube through a fine mesh screen into a container filled with sea water. The desired intermediate size zooplankton are retained in the lower filtering tu be. Upon completion, the upper filtering tube is removed and zooplankton in the lower tube are transferred to the container with a pipette. A whole set is made with Plexiglass or Perspex (after Omori and Ikeda 1984, 'Methods in marine zooplankton ecology' (ISBN: 047/-80/070), John Wiley & Sons, Inc., reprinted by permission of John Wiley & Sons, Inc.). With regard to the time del ay between actual capture of zooplankters and subsequent measurements, there is a trade-ofI between stress and starvation (Capture stress! starvation, section 10.3.1). Specimens should be allowed to recover from the trauma of capture for a period of time but not for so long that the metabolism drops significantly due to starvation. DifIerent investigators use difIerent recovery periods varying from 2 h to greater than 24 h. A few things should be kept clearly in mind when deciding for oneself how long to allow animals to remain in the unnatural environment of the laboratory befo re using them in experiments. First, any manipulation of the experimental subject is a potential source of stress or excitement. If the investigator is concerned with accumulation of oxygen debt during capture, similar concern should be extended after the introduction into the respirometer. Second, the animal is never going RESPIRATION 493 to be healthier and happier than it was when swimming in the ocean. Even if it can be maintained in the laboratory and fed for days or weeks it is unlikely in the extreme that it is improving in health. Thus, in principIe, the sooner after capture that a metabolic rate is taken, the better the result is likely to be. 1 10.7 RESPIRATION 10.7.1 Oxygen consumption -Winkler titration (T. Ikeda) The method described below largely follows Omori and Ikeda (1984). In the sealed chamber method, bottles with and without experimental zooplankters are prepared simultaneously and the difference in concentrations of dissolved oxygen after a period of incubation is attributed to respiration of the zooplankters. This method has been the most popular since the original oxygen consumption measurements on Calanus finmarchicus by Marshall el al. (1935). The major advantage of the water bottle method is its simplicity in that it requires no specialized equipment. However, the method requires relatively long incubations, therefore starvation cannot be completely eliminated as a potential inftuence on the rate (Technical problems, section 10.3.1). Since the changes in oxygen consumption rate of zooplankton during the incubation period cannot be determined, the method assumes that the oxygen consumption rates of zooplankters are constant during the entire incubation periodo INCUBATION BOTTLES BOD bottles are ideal, but ordinary glass reagent bottles fitted with airtight lids may also be used. The capacity of each bottle should be measured prior to its use in an experiment. The capacity may be obtained from the difference in the weight of each bottle with lids before and after filling it with distilled water (1.00 g = 1.00 mI). Prior to the experiment bottles and pipettes for the transfer of specimens from one container to the other should be cleaned with acid solution (6NHCI), then rinsed well with pure water and dried. WASHING/EXPERIMENTAL SEA WATER Water sampled from the site from which zooplankters were obtained is used to wash and subsequentIy incubate the zooplankters. In order to remo ve other organisms and particles, water should be filtered through GF/C, GF/F or Nuclepore filters with equivalent pore-size (0.4 11m). To remove smaller oceanic bacteria, Nuclepore filters with 0.2 11m are best. Vacuum filtration usually reduces the dissolved oxygen content in water. Therefore, aeration of the water is necessary to readjust the oxygen content to near saturation level. Most experiments start at oxygen saturation. Special attention may be required for experimentation on animal s from low oxygen habitats. To remove particles from low-oxygen water without appreciable changes in oxygen content, a gentle filtration system (see Figure 10.9B) using a combination of a siphon system and funnel covered with fine mesh netting (10 or 20 11m mesh openings) may be used. Three typical washing procedures are illustrated in Figure 10.10 A-C. The best washing method depends on the character of the zooplankters and the experimental designo Generally, the washing procedure is not so critical for oxygen consumption rate measurements with oxygen saturated water, but it is a very critical source of error for 494 METABOLlSM A B Cut c:{¡ O ~/~_ -O Netting 2 SyringeT1 3 Q Pipette ( lffij\ f U~.~ I t~~ tA~~,'.- ~ ~aker 4 ( ~ /2~._-:~~_ Filtered sea water \ (g ~:::::.. Experimental I ~!i"-- L.:J., I L1. bottle Petri dish D 2 3 Fig. 10.10 Three procedures for washing zooplankton for metabolic experiments, and preparation of bottles for incubation after washing. (A) A siphon system, originally designed by Marshall et al. (1935) for oxygen consumption measurements. In their original method, sea water volume needed for flushing is 6-7 times the bottle volume. (B) Transparent plastic syringe. Place animals in the syringe with a pipette, and rinse them in several beakers. Then, place the syringe into a petri-dish and introduce the animals into bottle with pipette. A convenient size of syringe is 20 mi (2 cm in diameter), but larger ones can be used when fine netting is placed over the tipo (C) Glass beaker for relatively large zooplankton. Gently add filtered sea water in the beaker and decant carefully. Repeat this treatment 3 to 4 times, and then place the animals into a bottle. (O) Preparation of a bottle for incubation. See text (after Omori and Ikeda 1984, 'Methods in marine zooplankton ecology' (ISBN: 047/-80/070), John Wiley & Sons, Inc., reprinted by permission of John Wiley & Sons, Inc.). RESPIRATION 495 ammonia and phosphorus excretion measurements mentioned in section 10.8. It is very important not to damage the specimens while washing them. The number of zooplankters should be counted before incubation (and again at the end of incubation). The bottles are filled with experimental water using a siphon system, placing one end of the siphon onto the bottom of the bottles. After zooplankters have been washed and placed into the bottles, lids should be fitted so as to avoid trapping any air bubbles. The lids are firmly wrapped with a plastic sheet and rubber band to reduce the risk of introduction of air bubbles in the bottles during incubation. Control bottles without zooplankters should be prepared concurrently using exactly the same procedure. In a typical experiment with ten experimental bottles, two control bottles are prepared befo re the first experimental bottle and two after the last experimental bottle. An additional control bottle may be inserted between the fifth and sixth experimental bottles (15 bottles in total). The bottles are ready for incubation after wrapping with aluminum foil or black plastic sheet (Figure 10.10D). A water bath with a temperature control unit is best for incubating the bottles. INCUBATION The incubation period will vary depending on the experimental temperature, density of zooplankters relative to the volume of the bottle, and rates of oxygen consumption of the zooplankters. For medium to large sized zooplankters, 12 or 24 h is recommended to eliminate the effect of diurnal rhythm on the metabolic rates. The period may be shortened to only a few hours for very active zooplankters that are less tolerant to prolonged starvation during the incubation periodo The length of incubation period should first be based on the size of zooplankton (i.e. tolerance to starvation). Decision on the size of bottles used in the incubation and the number of specimens in each should be based on the need to minimize accumulation of excreta and oxygen depletion at the end of the incubation (see Oxygen saturation, section 10.3.1). Antibiotics, such as streptomycin and chloromycetin, are often used to minimize bacterial activity in the water. The difficulty with using antibiotics is the determination of dosages, which vary among different zooplankters. Unfortunately, líttle is known about the appropriate dose of antibiotics for different zooplankton, and excess antibiotics may seriously affect the activity of experimental animals. Sorne antibiotics may interfere with other chemical analyses (Bacteria, section 10.3.1). To determine dissolved oxygen, a volume of water is removed from the bottle after incubation. Figure 10.11 shows a siphon system that we use to transfer water from incubation bottles into small oxygen bottles. Duplicate water samples are sufficient when the analyst is skilled in the technique. At the end of the incubation, the activity of zooplankters in the experimental bottles should be checked. These zooplankters may then be transferred to a petri-dish (larger zooplankters) or directly onto a piece of mesh (smaller zooplankters). Specimens can be counted either with the unaided eye or, for smaller zooplankters under a dissecting microscope. Zooplankters should then be weighed (wet weight) and/or weighed after drying (dry weight). Prior to weighing, water adhering to the body of a specimen should be removed by placing the specimen on filter paper. For smaller zooplankters that are difficult to han dIe individually, this can be achieved by placing the filter paper on the other side of the mesh. Although a brief rinse with distilled water to remove salts is desirable it is sometimes accompanied by loss of integrity, and organic matter, due to body lysis from osmotic shock (see Omori 1978). 502 METABOllSM the electro de will no longer be depleted in oxygen by the consumption of the electrode itself. In this manner, an electro de with a larger cathode can be used without the need for stirring. There is no reading taken during the time the electro de is off between pulses, making this design a liule reminiscent of sequential sampling using the Winkler technique. Nevertheless, the electrodes employing this circuit design are widely used and have a good reputation for accuracy. CHAMBERS Apart from three basic requirements, the design of chambers for sealed jar respirometry is limited only by the needs and creativity of the investigator. The three basic requirements are, first, that the chamber be able to accept the electrode tip; second, that the chamber-electrode combination are sealed to the atmosphere; and third, that there is sorne means for rigorously controlling temperature within the chamber. Two us~ful chamber designs are shown in Figures 10.14 and 10.15 (cf. Quetin and Mickel 1983). In Figure 10.14 the tip of a syringe is cut off and the electro de itself forms an opposing plunger to the syringe plunger. In Figure 10.12, a water-jacketed lucite chamber has a chimney that mates with an O-ring on the barrel of the electrode. The syringe barrel chamber can be submerged within a water bath for temperature control while the water-jacketed chamber is connected to a circulating refrigerated water bath. The design shown in Figure 10.14 has been successfully employed for respiratory determinations on individual copepods and fish larvae as small as 2 mm using unstirred electrodes. The design in Figure 10.15 is more suitable for larger species such as euphausiids and amphipods and can be used with stirred or unstirred electrodes. The cut-off for the minimum size of animal that is amenable to being an experimental subject Plunger Oxygen electrode Fig.10.14 A microrespirometer using an oxygen electrode and a cut syringe. The same O-ring that seats the electrode membrane seals the respirometer to the atmosphere. The assembled syringe-electrode combination as shown is placed in a water bath for temperature control. Alternatively, one can use the same principie with a luer-Iock valve on the end of the syringe and using the electrode in place of the syringe plunger. The valve obviates the need for cutting the syringe. Chamber Water-retaining lip Water jacket Vent to release water displaced by electrode RESPIRATION 503 Stainless steel bolts Inlet to water jacket Fig.10.15 A water-jacketed chamber for measuring oxygen consumption in larger zooplankton. The O-ring midway up the oxygen electrode makes a gas-tight seal on the walls of the chimney. A water-retaining lip allows the chamber to be overfilled to aid in eliminating bubbles. The chamber lid seats on an O-ring seal and is held down with wing nuts. The entire chamber is manufactured from lucite. Design by Joe Donnelly. with stirred electrodes must depend on the goodjudgment ofthe investigator. Clearly, if an animal is swimming hard to maintain a position within the respiratory chamber against the current created by the stir bar, stirred electro des are not appropriate. In no case is it appropriate to use stirred electro des with gelatinous species. Most investigators will be using electrodes available from commercial sources. Many come with choices of chamber types that are designed to mate with their electrode; sorne are very 10w vo1ume systems that are suitab1e for use with zooplankton. If the investigator has just spent a considerable sum of money on an electro de system, he or she will be loathe to put their electrode in a lathe to modify it. In those cases the creativity and machine shop work will have to be restricted to the respiratory chamber. The investigator should, where possible, opt for flexibility in their chosen system. PITFALLS Temperature is a critical factor in respiratory determinations with the oxygen electrode, not only because of the effect of temperature on the metabolism of ectotherms but because oxygen electrodes are very sensitive to temperature. In fact, electro des are more sensitive to tempera tu re than they are to oxygen because the electrochemical dissociation occurring at the electro de tip is governed by the same QlO rule as any biochemical reaction. Thus, while the solubility of oxygen at 10 oC is 123% ofthat at 20 oC, the current 504 METABOLlSM flow generated at the electrode tip at 10°C will be about 50% and so will the electrode output. Sorne commercially available electrodes are available with temperature compensation but keeping experimental and calibration temperatures identical is the simplest and best procedure. When using the small volumes typical of experiments with individual zooplankton, bubbles can be a large potential problem in respiratory determinations. Oxygen is about 30 times more concentrated in air than in sea water; a very small volume of air can perturb the accuracy of a respiratory determination made in, for example, 2 mI of sea water. The investigator must be meticulous in removing all bubbles from the respiratory chamber. The best way to prevent bubbles is to avoid introducing them into the chamber to begin with. Siphoning water, using a large syringe, or pouring gently into your respiratory chamber will prevent introduction of bubbles. Once bubbles are in, microbubbles can be chased with larger bubbles or brushed out with small artist's brushes. Do not introduce cold water into a respiratory chamber and allow it to warm up. Bubbles will form as the solution degasses due to the reduced solubility of gases at warmer temperatures. Allow the water to equilibrate to experimental temperature first. EXPERIMENTAL PROTOCOL Electrodes are best calibrated immediately prior to, and directly after, experiments. In most cases, a high calibration is obtained by gentle aeration of the electrode in a calibration bath at experimental temperature or by using a combination of stirring and aeration. A low calibration is obtained by using a more violent bubbling with nitrogen gas in the calibration bath to drive off the oxygen in solution. Sodium sulphite is sometimes used to obtain a low calibration, but there is potential for poisoning a platinum cathode (see the five-step reaction given aboye) and it should be used with caution. Nitrogen gas, though inconvenient, does nothing but purge the oxygen from solution. Electrodes should ideally come to within 3 % of electrical zero at the low end. Electrodes should be stable at the low and high ends and there should be little or no difference in high and low calibrations before and after the runo Differences of greater than 5% in the preand post-run high calibrations constitute grounds for rejection of a runo Linearity is assumed between the low and high points on the calibration. The sea water to be used as a respiratory medium should be well filtered to remo ve microorganisms. The author (J.J. Torres) uses membrane filters (0.45 11m pore size) and has had little problem with control rates. Experimental subjects should be transferred to filtered water at experimental tempera tu re prior to introduction into the respirometer as a rinsing step. After a satisfactory calibration has been obtained, the experimental subject is introduced to a bubble-free respiratory chamber and, taking extreme care not to introduce bubbles or disturb the subject more than absolutely necessary, the chamber is sealed, the electrode inserted, and the run is allowed to begin. It is best if visual stimulation is kept to a minimum by shading or isolating the animal in sorne way. If stirring is to be used, insuring that the magnetic stirring motor is fully warmed up prior to the run will prevent the magnetic stirrer speed-up that can result in homogenization of your experimental animal. It is best to avoid stirrer-hotplate combinations because both malfunctions in the hotplate and oversights due to fatigue can result in an unintended bouillabaisse instead of an experiment. Control runs are used mainly to check for contributions by microorganisms to the total measured oxygen consumption rate. Since most microorganisms will be introduced by the experimental subject, it is best to take control rates at the termination of a run by removing the subject, replacing its volume with fresh filtered sea water, and RESPIRATION 505 re-starting the runo Keeping respiratory chambers c1ean and using only filtered sea water for electro de calibrations and respiratory runs will go a long way to preventing errors when measuring control rates. Other uses for control runs are to check for electrode drift and oxygen consumption by the respiratory chamber or respiratory medium. In the latter two instances runs would be set up as normal, but without an animal. Antibiotics are sometimes added to the filtered sea water employed as a respiratory medium to prevent microbial growth in the respiratory chamber. Those most commonly used are penicillin, streptomycin, and neomycin at concentrations of 25 to 50 mg 1 - l. The authors' (J.J. Torres) experience has been that microbial growth is a negligible problem within the time frame of a typical run (8 to 12 h) when working with filtered sea water at temperatures less than 15 oC. Microbial growth needs to be more carefully monitored at higher temperatures and antibiotics can help to control this. Runs greater than 12 h in duration need to be checked carefully for microbial contributions at any temperature but especially at temperatures aboye 15 oC. The recording apparatus should be consulted frequently. Notes on animal activity and condition should be made at intervals as unobtrusively as possible. Low oxygen becomes stressful to most pelagic species at about 25% of air saturation (e.g. Donnelly and Torres 1988; Torres el al. 1994); they can no longer maintain a constant oxygen consumption rate below this leve!. It is best if the chamber volume allows the animals to remove 5% to 100/0 of the oxygen in the chamber each hour. CALCULATIONS AND TREATMENT OF DATA Calculations of metabolism with data from oxygen electrodes use the same principIes regardless of the recording system. Two leaps of faith are required when equating the electrical output of oxygen electrodes to quantities of dissolved oxygen. The first is that the electrical current produced by the electro de varies directly and linearIy with the dissolved oxygen in the range from zero to air saturation. The second is that the high calibration point is equivalent to the oxygen solubility at the temperature and salinity in the calibration bath. The assumption oflinearity can be checked by testing the output ofthe electrode at air saturation as you would during a normal calibration and then at oxygen saturation by using a tank of pure oxygen. This will then give you the output at 0%, 21 % and 100% O2, enough points to check for linearity of response. Alternatively, you can use the Winkler method or a gas chromatograph to check at various points between zero and air saturation. The author (J.J. Torres) has never noted a problem with linearity. The assumption that the high calibration point of the electrode is equivalent to the oxygen solubility at the calibration tempera tu re and salinity is necessary to convert the electrical output of the electrode to the units of dissolved oxygen that are of use to USo Values for oxygen solubilities over a wide range of temperatures and salinities are available from the exacting research of marine chemists. The most widely accepted numbers for oxygen solubility are those from the equation of Weiss (1970), who, using the data of Carpenter (1966) and Murray and Riley (1969), described the relation between temperature, salinity and oxygen solubility. Weiss's equation, as reported in Kester (1975; equation 8.11) is: lnc* = Al + A2(100/T) + A3In(T/100) + A4(T/100) + SO/oo[B I + B2 (T /100) + B3(T /100)2] (10.16) 506 METABOLlSM for c* = O2 solubility in ~mol kgl based on the data ofCarpenter (1966) then: Al = -173.9894, A2 = 255.5907, A3 = 146.4813, A4 = -22.2040, BI = -0.037362, B2 = 0.016504, and B3 = -0.0020564, and for c* = O2 solubility in cm3 l1 based on the data of Murray and Riley (1969) then: Al = -173.4292, A2 = 249.6339, A3 = 143.3483, A4 = -21.8492, B1 = -0.033096, B2 = 0.014259, and B3 = -0.0017000. Values as sume an atmosphere of20.94% O2 at 760 mmHg total pressure and at 100% relative humidity. As an alternative to using the equations above, one can look up the tabulated values in either of the original papers and use those. The ditTerences in solubilities between the original papers and Weiss's improved numbers are small enough (~ 0.03 cm3 dm - 3) for our purposes to make the error an acceptable one. Reading any of the original papers on oxygen solubility and especially Kester's treatment will familiarize the investigator with units, accuracy, and precision in determining values for dissolved gases. The electrode output at the low and high calibration points is assumed to be 0% and 100%of air saturation respectively. The O2 solubility determined from Weiss's equations or from the original tabulated data at the experimental temperature and salinity yields the total dissolved oxygen at air saturation. The investigator can then compute the oxygen consumption from the percent change in total electrode output with time. For example: • electrode output at high cal is 100 mV which equals O2 solubility at air saturation: 7.10 mll1 • electrode output at low cal is O m V • total volume of respirometer: 5 mI • total oxygen in system: 35.5 ~l • wet mass of experimental subject: 10 mg After 1 h the electrode reads 90 m V, 10% of oxygen in the system has been removed. Oxygen removed from system is 3.55 ~1, oxygen consumption rate is 0.355 ~l O2 mg wet mass -1 h1. 110.7.3 Enzymatic method -electron transfer system (S. Hernández-León) The Electron Transfer System (ETS) is the pathway responsible for transfer of electrons to oxygen, the final electron acceptor. As a consequence, its activity is responsible for oxygen consumption by both the cell and organism, and can be used as an index, or biochemical proxy for zooplankton respiration in the sea (section 10.4.1). ETS may be characterized as a multi-enzyme, multi-substrate system, and its activity is determined in substrate saturating conditions, i.e. at the maximal rate (V max )' The substrates are nicotinamide adenine dinucleotide (NADH), nicotinamide adenine dinucleotide phosphate (NADPH), and succinate, which satura te microsomal and mitochondrial ETS. The tetrazolium salt 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride (INT) is used as an artificial electron acceptor to measure the electron transmission rate: a mole of oxygen consumed will be equivalent to two moles of INT reduction. The slight1y yellow INT is reduced to the pink colored formazan and the intensity of the color is read with a spectrophotometer. The enzymes in volved in the electro n transfer system are NADH dehydrogenase (EC 1.6.99.3), NADPH dehydrogenase, (EC 1.6.99.6), NADPH-cytochrome reductase (EC 1.6.2.4) and succinate dehydrogenase (EC 1.3.99.1). The activity of these enzymes are recorded in the same assay and the ETS activity is not necessarily the addition of the RESPIRATION 507 activities ofthe different enzymes of an enzyme complex (Savenkoff et al. 1995; Gómez et al. 1996). NADH dehydrogenase is the most active dehydrogenase in zooplankton material. Since the work by Packard (1971) the assay method has been modified by the addition of Triton X-lOO (Owens and King 1975), and by changes in pH of the buffers (Kenner and Ahmed 1975). The assay method described here is essentially the same as that of Kenner and Ahmed (1975) and Packard and Williams (1981), modified slight1y by Gómez et al. (1996) for zooplankton. REAGENTS 1) Homogenization buffer 0.05 mol phosphate buffer, pH 8.0, containing 6.72 g Na2HP04, 0.362 g KH 2 P0 4, 1.5 g polyvinyl pyrrolidone (PVP) , 18.5 mg MgS0 4.7H20 and 2 mI Triton X-lOO, in 11 of deionized-distilled water (DDW). Store at - 20 oC. 2) Daily homogenization buffer Add, immediately before use, an appropriate amount ofO.1 mol NaCN (e.g. 0.735 g NaCN in 150 mI ofDDW) to obtain a final concentration of 2 mmol NaCN (e.g. 1 mI NaCN solution in 49 mI of the homogenization buffer). 3) Substrate buffer Prepare a solution ofO.05 mol phosphate buffer, pH 8.0, containing 6.72 g Na2HP04, 0.362 g KH 2 P0 4, 36 g disodium succinate hexahydrate and 2 mI Triton X-lOO. Store at - 20 oC. 4) Daily substrate solution Just before the assay, dissolve 15 mg NADH and 5 mg NADPH in 24 mI ofthe substrate buffer. 5) Blank substrate buffer Prepare the substrate buffer (3) without substrates as a blank. Store at - 20 oC. 6) INT solution Prepare a solution of the tetrazolium salt 2-(4-iodophenyl)-3-(4nitrophenyl)-5-phenyltetrasolium chloride (lNT) in DDW up to a concentration of 4 mmol (e.g. 1 g INT in 500 mI DDW). Filter the solution and store at - 20 oC in the dark. 7) Quench solution 50% formalin (36%) plus 50% 1 mol H3 P0 4, pH 2.5. Store under refrigeration (0-4 OC). Bacterial contamination should be kept to the absolute minimum when mixing up solutions. To achieve this, working in very c1ean conditions is essential, and solutions should not be kept for long at room temperature. SAMPLE PREPARATION ANO PRESERVATION Sorted live zooplankton is ready to use for the assay after removing excess water with blotting paper. For mixed zooplankton, samples in the net cod-end are poured onto a 100 f.1m mesh sieve and washed with filtered sea water. The mesh sieve can then be put over blotting paper to remove excess water. If samples are too large to apply this procedure, they may be gent1y vacuum filtered. Samples are scraped offthe filters using a spatula for the assay. Fresh samples are best, but freezing samples may be inevitable in the experiments at sea. Ahmed et al. (1976) observed that although a cell-free extract ofthe copepod Calanus finmarchicus frozen at - 2 oC lost considerable ETS activity in 24 h, no measurable loss was found when intact specimens were stored at - 20 oC or be10w for at least one week. No significant loss of ETS activity during storage for 34 days at - 20 oC has been reported for two fish species (lkeda 1989b). On the other hand Bamstedt (1980) found 508 METABOLlSM that frozen intact specimens of Acartia tonsa 10st 50% of their ETS activity within 30 h at - 20 oC and virtually no activity was detected after 72 h. Recently, Gómez et al. (1996) observed no significant 10ss in ETS activity on zooplankton samples preserved in liquid nitro gen (- 196 OC) for at least 2 months. Therefore, preservation of zooplankton samples in liquid nitrogen is recommended for the ETS assay, whenever the immediate assay is not practicable. ASSA y PROCEDURE 1) The zooplankton sample is homogenized with or without GF /C filters (Notes for the assay procedure and calculations, section 10.7.3, part a) in a teflon-glass tissue grinder (Notes for the assay procedure and calculations, 10.7.3, part b) for 2 min at Oto 4 oC in a known volume ofhomogenization buffer (approximately 10-20% (v/v) homogenate) (Notes for the assay procedure and calculations, section 10.7.3 part c) 2) The homogenate is then poured into a chilled graduated cylinder in order to record the vo1ume. The difference between this volume and the volume of buffer used for homogenization is the wet volume of the sample. 3) Centrifuge (Notes for the assay procedure and calculations, section 10.7.3 part d) for 10 min at 1000 g at O to 4 oC. Because the method is very sensitive the supernatant must be diluted in homogenization buffer. A 3 to 61 x dilution is often used depending on the concentration of sample in the homogenate. 4) Take al-mI aliquot of diluted supernatant, mix with 3 mI of the daily substrate solution and 1 mI ofINT solution, and incubate for 20 min at the in situ temperature in the dark. The color of the reaction mixture changes during the incubation period to pink due to the reduction of the INT to formazan. Red color is indicative of excess sample. Concurrently, another 1 mI ofthe diluted supernatant is incubated in the blank substrate buffer (substrate blank). Get another blank to account for the reaction between substrate and INT (reagent blank). Take 1 mI of daily homogenization buffer and incubate using the same procedure as with the sample. 5) Stop the reaction with 1 mI of the quench solution. 6) Read absorbance with a spectrophotometer at 490 nm and at 750 nm as the turbidity base lineo The increase in absorbance can be monitored continuously by using a temperature-controlled cuvette placed in a spectrophotometer. New technologies such as microplate readers and robotic samplers can also be used to save time when a large number of samples has to be processed. In order to obtain specific rates, measure protein contents of the homogenized samples obtained at step 3, by the method of Lowry et al. (1951) using bovine serum albumin as the standard, or modified Lowry et al.'s method by Peterson (1977, 1983) for samples with very low protein content. Please note that Savqenkoff et al. (1995) suggested the use of a yeast (Saccharomyces cerevisiae) as a reference to standardize ETS assays. The yeast is available in dried form which can be weighed and stored. The procedure consists of weighing five samples of the yeast from 1 to 5 mg with an ultramicrobalance using its large grains. Follow the steps given in the assay procedure but homogenize together with GF/C filters (diameter: 25 mm) in order to disrupt mitochondria walls. Plot weights against ETS activity. ETS activity data from dissimilar assay procedures by different workers could be compared with the reference material. CALCULATIONS ETS activity is calculated as: ETS = 60 x H x AS x COD/(INT x T x L xf), COD = [(AOD x AS) - (ROD x RS) - (FOD x FS)]/AS, RESPIRATION 509 (10.17) (10.18) where H is the homogenization volume, AS is the assay volume, BS is the blank volume, FS is the reagent blank volume, AOD is the difference between the spectrophotometer readings at 490 and 750 nm of the assay (BOD and FOD are the same differences for substrate blank and reagent blank, respectively), Tis the incubation time (minutes),jis the volume of homogenate used in the assay (ml), L is the path length of the spectrophotometer cuvette cell used (usually 1 cm), 60 is the factor to convert minutes to hours, and INT is the standardization factor (see below) which converts the absorbance of the produced INT -formazan. The ETS activity is given in fIl 02h -1 but could be expressed in electron equivalents (fIEq e -) by dividing the ETS activity by 5.6 (1 mol Of02 = 4 Eq eand ETS x 4/22.4 = ETS/5.6). ETS assays are normally run at one temperature during a set of experiments, or during a cruise. The experimental temperature may not be the in situ temperature. ETS activity obtained at a given incubation temperature may be converted to the activity at in situ temperature using the following equation: ETS in silu = ETS incu X exp[E a (l/ Tincu - 1/ T in silu)/ R] The value of the Arrhenius activation energy (Ea) is about 15 kcal mol1 in Calanus finmarchicus and in the range of 13 to 16 kcal mol1 for a variety of other plankton. The symbol R is the gas constant (l.987 x 103 kcal mol1) and T is the absolute temperature (Packard el al. 1975). Convert ETS activity to respiratory oxygen consumption rate using the correlation between respiration rate and ETS activity (R/ ETS ratio). The R/ ETS ratio is theoretically around 0.5. Problems associated with the R/ ETS ratio are detailed in ETS activity, section 10.4.l. INT STANOAROIZATION Because of different qualities (or different trade marks) ofINT, it is highly recommended that you standardize it using the procedure given below. REAGENTS ANO SOLUTIONS 1) 0.1 mol phosphate buffer Add 13.6 g KH 2 P0 4 to 11 of DDW. Add 14.2 g Na2HP04 to 11 of DDW. Mix 808 mI of the Na2HP04 solution with 192 mI of the KH 2 P0 4 solution. Add 10 mI Triton X-lOO to the mixed solution. Store at -20°C. 2) INT color reagent Add 8 mg phenazine methosulphate (PMS) and 0.4 mI Triton X-lOO to 16 mI ofthe INT solution to be standardized. 3) 0.05 mol phthalate buffer Dissolve 2.55 g phthalic acid in 172 mI of DDW. Add 51 mI ofO.1 NHCl and 22 mI Triton X-lOO. Adjust to pH 3.0 ifnecessary. Dilute to 250 mI. Standardization procedure 1) Prepare a solution of 0.043 fImol ml1 of NADH in 0.1 mol phosphate buffer, pH 7.4 (e.g. 2 mg NADH in 60 mI ofO.1 mol phosphate buffer). 2) Prepare a series of dilutions to obtain different concentrations between O and 0.258 fIEq e-in a total volume of 3.0 mI. 510 METABOLlSM 3) Add 2.0 mI of INT color reagent to each tube and incubate in darkness for exact1y 1 min at room temperature. Stop the reaction with 1.0 mI of 0.05 mol phthalate buffer. Total volume ofthe assay is 6.0 mI. Read absorbance at 490 and 750 nm with a spectrophotometer. The factor used in the standardization is calculated converting the ,umol NADH mll to ,uEq e - (multiply ,umol mll by 2). Plot absorbance against the ,uEq e - and obtain a slope from a least square linear regression. Then, multiply the slope by 1.24 obtaining the value to convert the absorbance of the INT-formazan produced in the calculation of ETS. This value should be around 1.42 because the INT -formazan has a molar extinction coefficient (A 490 ) in 0.133% Triton X-lOO solution of 15.9 x 10 3 moll cm -l. INT requires two electrons to be reduced to formazan and oxygen requires four electrons to be reduced to water. Therefore, the extinction coefficient ofINT-formazan is equivalent to a 0.5 molar solution or 11.2 1 O2 1 -1. Because the solvent volume is taken into account in the ETS assay, the equivalent absorptivity of 1 ,u1 O2 per mI of solvent willbe 15.9 x 10 3/(11.2 x 10 3) = 1.42. NOTES FOR THE ASSAY PROCEOURE ANO CALCULATIONS a) The use of GF/C filters is to facilita te disruption of mitochondria walls. However, Gómez el al. (1996) found no significant difference between ETS activities of mixed zooplankton samples processed with and without GF/C filters, and suggested that chitin exoskeleton of crustacean zooplankton had the same function as GF /C filters for crustacean zooplankton samples or mixed zooplankton samples which inelude crustaceans. b) A Polytron ultrarapid homogenizer may be used instead, as Gómez el al. (1996) noted no significant differences between the resultant ETS activities of samples processed with teflon-glass grinder and the Polytron ultrarapid homogenizer. Sonication may also be an alternative, but the author (S. Hernández-León) has not tested this yet. c) Crude homogenates of zooplankton can be maintained in a water-ice bath (0-4 OC) for up to 90 min without any significant loss in ETS activity (Bamstedt 1980; Gómez el al. 1996) thus allowing preparation of several samples before the start ofthe assay. d) While Gómez el al. (1996) observed no significant differences in ETS activities of samples centrifuged before and after the incubation, the procedure given here (centrifuge before the incubation) is recommended to avoid an increment of turbidity in the assay. 1 10.7.4 Enzymatic method -Iactate dehydrogenase and citrate synthase (11 Torres and S.P. Geiger) Lactate dehydrogenase (LDH) and citrate synthase (CS) catalyze key reactions of anaerobic and aerobic intermediary metabolism respectively. LDH is the terminal enzyme of anaerobic glycolysis as the pathway is conventionalIy presented; it catalyzes the conversion of pyruvate to lactate and is one of the three rate-limiting steps in the pathway. CS catalyzes the condensation reaction between acetyl-CoA and oxaloacetate to form citrate, which is the first and rate-limiting step in the 'Krebs cyele. The rationale behind using LDH and CS as indicators of metabolism is, first, that the activity of each enzyme is indicative of the metabolic poise (cf. Hochachka and Somero 1984), or activity, of its respective pathway, and second, that the activity of the pathway relates RESPIRATION 511 directly to the metabolism and therefore the overall physiological condition of the whole organism (see Enzymes ofintermediary metabolism in section 10.4.2). LDH is best applied as a physiological indicator in ichthyoplankton (Clarke et al. 1992; Brightman 1993). The data available suggest that LDH activities in ichthyoplankton correlate well with growth and physiological condition, and are further supported by years of research on the biochemical physiology of vertebrates (Hochachka and Somero 1984). Less is known about the function ofLDH in crustaceans and other zooplankton. It is present in detectable quantities (e.g. Thuesen and Childress 1993a, 1993b, 1994; Geiger et al. 1996; Thuesen et al. in press) but its function in invertebrate metabolism is less well described and variability even between species of a general taxonomic group, e.g. calanoid copepods, can be very high. For example, results on LDH activities in Antarctic copepods (S.P. Geiger and J.J. Torres unpublished data) using the techniques described in Lactate dehydrogenase, section 10.7.4 suggest low, but detectable, levels of LDH, whereas results described in Thuesen et al. (in press) suggest very high activities in deep-living copepods of the California Current. Invertebrates exhibit considerably more creativity with their glycolytic endpoints than do vertebrates (Hochachka and Somero 1984). Thus, the meaning of LDH activity should be evaluated for invertebrate preparations on a case-by-case basis. CS, while it is a slightly more difficult assay, seems to work equally well on both vertebrate and invertebrate subjects and is especially useful in copepods. We recommend it as a first choice for assaying condition in copepods and LDH as a first choice in ichthyoplankton. TISSUE PREPARATION ANO GENERAL PROCEOURES Enzyme activities are most stable when tissue is frozen initially in liquid nitrogen, and then stored at - 80 oC. No loss in enzyme activity occurs for at least several months and for most enzymes, storage at - 80 oC allows nearIy indefinite storage. Both LDH and CS are fairIy stable enzymes and if the ideal situation cannot be achieved, then investigators should do their best to approach them and be wary of deterioration. If a choice must be made, the initial (liquid nitrogen) freezing step is the most critical one and the samples can usually then be stored in a conventional (- 20 OC) deep-freeze with little loss of activity for a few months. A word of caution: tissues and enzymes do vary in their sensitivity to storage, so these procedures should be viewed as general guidelines and not gospel. Gelatinous organisms and small copepods are particularIy sensitive to desiccation with long term storage. It is best to determine empirically the sensitivity of your own preparations to storage. Once homogenized, samples should be used within several hours and not be refrozen. Homogenize tissue in ice-cold 50 mmol imidazole buffer (pH 7.2 at 20 OC) using either a sonicator or conventional homogenizer. The dilution factor to be used varies with the tissue and its inherent enzyme activity. It is imperative that the dilution factor be known precisely because it is an important element in calculating enzyme activity and expressing it in standard units (see Calculations, section 10.7.4). For tissues with low activity, a dilution factor of 1 to 4 (l g tissue to 4 mI buffer) is useful, for tissues with more enzyme a 1 to 24 dilution is more appropriate. Large tissue samples (e.g. Euphausia superba abdominal musc1e) should be ground in chilled ground glass hOl1)ogenizers, and then briefiy sonicated. For small tissue samples (e.g. calanoid copepods, or young < 10 mm fish larvae) sonications usually provide adequate homogenization. Large tissue samples are best centrifuged at 2500 to 5000 g for 10 min to reduce particulate interference during the assay. 518 METABOLlSM AMMONIA-N Reagents 1) Deionized water Remove the ammonia from distilled water by passing it through a small coIumn of cation exchange resin in the hydrogen form just before use and store the water in a tightly stoppered glass flask. 2) PhenoI soIution Dissolve 20 g of crystalline phenoI (analytical reagent grade) in 200 mI of95% (v/v) ethanol. 3) Sodium nitroprusside solution Dissolve 1.0 g of sodium nitroprusside Na2Fe(CNhNO·2H20 in 200 mI of deionized water. The solution is stored in a brown bottle. It is stable for about a month. 4) Alkaline solution DissoIve 100 g of trisodium citrate and 5 g of sodium hydroxide (analytical reagent grade) in 500 mI of deionized water. This solution is stable for a long periodo 5) Sodium hypochlorite solution Use a solution of commercial hypochlorite (e.g. Chlorox) which should be at least 1.5N (see Note a). 6) Oxidizing solution Mix 100 mI ofreagent (4) and 25 mI ofreagent (5). It is best to prepare this solution immediately befo re the analysis. Analytical procedure Add 10 mI of the sample water (see Note b) to a test tube with an accompanying screw cap and then 0.4 mI, 0.4 mI and 1 mI of soIutions (2), (3) and (6) respectively. Mix well after each addition. Fit the cap to the test tube in order to avoid contamination from ammonia in the air and allow to stand at a temperature between 20 to 27 oC for 1 h. Then measure the extinction at 640 nm relative to distílled water in a spectrophotometer using lO-cm cells. It is best to conduct the reaction in a constant-temperature water bath. The reaction requires a full 60 min for completion. During that time the samples should never be placed in direct sunlight or near a window. The detrimental effect of sunlight on the reaction has been pointed out (Liddicoat el al. 1975). Ammonia-N concentration of the sample water can be obtained from the following equation (see Note c): tlg-at N 11 = (E -Eh) x F (10.24) where E is the extinction of the sample water (a mean of duplicate readings), Eh is the extinction of reagent blank, and F is the factor. For Eh and F, see Calibration, section 10.8.3. 1 tlg-at N is equivalent to 14 tlg N. Calibration Dissolve 0.6607 g of ammonia sulfate (analytical reagent grade) in 11 of deionized water (1 mI == 10 tlg-at N). Add 1 mI of chloroform and store in a dark place with a stopper. At the time of use dilute the standard solution 100 times with distilled water to make a secondary solution, and dilute this solution further with filtered sea water containing as little ammonia as possible to make the standard solution. If sea water is added to 10 mI of this secondary solution to make 1 1 of the standard solution, the resulting ammonia concentration is equivalent to 1.0 tlg-at N 11 of ammonia-N. Pipette 10 mI of diIute standard into each of three test tubes and carry out the ammonia determination described in Analytical procedure, section 10.8.2. Calculate the factor, F, as where Estd is the average extinction of three standards and Eh is the average extinction of EXCRETION 519 the reagent blank. To obtain Eh, use deionized water in place of dilute standard solution and carry out ammonia determination. NOTES FOR THE AMMONIA ANALYSIS a) To check on the strength of hypochlorite, dissolve 12.5 g of sodium thiosuIfate (Na2S203·5H20) in 500 mI of distilled water. Add a few crystals (ca. 2 g) of potassium iodide (KI) to about 50 mI of distilled water in a small flask and pipette in 1.0 mI of hypochlorite solution. Add 5 to 10 drops of concentrated hydrochloric acid (HCl) and titrate the liberated iodine with the thiosulfate solution until no yellow color remains. Discard the hypochlorite when less than 12 mI ofthiosulfate is used. b) Temporary storage of sea water prior to analysis appears satisfactory in glass or polyethylene bottles, but analysis should not be de1ayed for more than 1 to 2 h at the most. If the analysis cannot be performed in this time period, samples should either be frozen at - 15 oC or sto red unfrozen in the presence of 0.4 mI of phenol solution (Reagent (2)) per 10 mI of sample. Samples may be stored in either manner for up to 2 weeks (Degobbis 1973). c) It assumes that the extinction is linear over the range of ammonia concentrations analyzed. For a linearity cheek of extinctions against ammonia concentration, a serial dilution of the standard solution should be made once during the experiment. The range of extinctions should cover the expected range of readings of experimental values. INORGANIC PHOSPHATE-P Reagents 1) Ammonium molybdate solution Dissolve 15 g of ammonium paramolybdate (NH 4) M0 70 24 ·4H20 (analytical reagent grade, preferably fine crystals) in 500 mI of distilled water. It should be protected from direct sunlight and be preserved in a polyethylene bottle. This solution is stable indefinitely. 2) Sulfuric acid solution Add 140 mI of concentrated sulfuric acid (analytical reagent grade, specific gravity of 1.8) to 900 mI of distilled water. Allow the solution to cool and preserve in a glass bottle. 3) Ascorbic acid solution Dissolve 27 g of good quality ascorbic acid in 500 mI of distilled water. Store the solution frozen in a poIyethyIene bottle. Thaw for use and refreeze at once. This solution can be kept for onIy about 1 week at room temperature but it is stable for several months if frozen. 4) Potassium antimonyl-tartrate solution Dissolve 0.34 g of good quaIity potassium antimonyl-tartrate C2H2(OHhCOOKCOO(SbO)·1¡2H20 in 250 mI of warm distilled water. The soIution shouId be preserved either in a gIass or polyethylene bottIe. It is stable for several months. Analytical procedure Immediately before the analysis, the aboye soIutions (1), (2), (3) and (4) are mixed in the ratio of2:5:2:1 (v/v) respectively. Use this reagent for one-batch sampIes and discard any excess; it should not be kept for more than 6 h. To 10 mI of sample water (see Note a) in a test tube, add 1 mI of the mixed reagent and mix immediateIy. After 5 min or at most within 1 to 2 h, measure the extinction of the soIution reIative to distilled water in a 10cm cell at a wavelength of 885 nm. 520 METABOLlSM Phosphate-P concentration of the sample water can be obtained from the following equation (see Note b): Jlg-at P 11 = (E -Eh) x F (10.25) where E is the average extinction of the sample water (a mean of duplicate readings), Eb is the average extinction of reagent blank, and F is the factor. For Eb and F, see Calibration, section 10.8.3. 1 Ilg - at P is equivalent to 31 Ilg P. Calibration Dissolve 0.816 g of anhydrous potassium dihydrogen phosphate KH 2 P0 4 in 1 1 of distilled water (1 mI == 6.0 Ilg-at P) and store in a dark bottle with 1 mI of chloroform. The solution is stable for many months. Dilute 10 mI ofthe standard solution to 11 with distilled water (1 mI == 6.0 x 102 Ilg-at P). Pipette 0.5 mI of dilute standard into each of three test tubes and make up to 10 mI with distilled water (3.0 Ilg-at P 11 ). Carry out the phosphate determination described in Analytical procedure, section 10.8.3. Calculate the factor, F, as (10.26) where Estd is the average extinction of three standards and Eb is the average extinction of the reagent blank. To obtain Eb use distilled water in place of dilute standard and carry out the phosphate determination. NOTES FOR THE INORGANIC PHOSPHATE ANALYSIS a) As the storage of sample water for inorganic phosphate analysis is always associated with uncertain errors, immediate analyses are recommended (Gilmartin 1967). b) It assumes the extinction is linear over the range of phosphate concentrations analyzed, and this is usually the case for phosphate analysis. For linearity check of extinctions against phosphate concentration, a serial dilution of the standard solution should be made once during the experiment. The range of extinctions should cover the expected range of readings of experimental values. 1 10.9 REFERENCES Aarset, A.V. and Aunaas, T., 1990. 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