EFFECTS OF AMMONIA ON GROWTH AND METABOLISM IN TILAPIA, OREOCHROMIS NILOTICUS By RICHARD JAMES MORROW A thesis submitted to the Department of Biology in conformity with the requirements for the degree of Master of Science Queen’s University Kingston, Ontario, Canada August, 2009 Copyright © Richard. J. Morrow, 2009
ABSTRACT Nile tilapia (Oreochromis niloticus) is an important species in the expansion of aquaculture, which supplements strained natural fish stocks worldwide. Although nitrogen accumulation in aquaculture has been documented as hazardous, recent studies have highlighted its potential to positively affect fish growth. The current study investigates the growth and oxygen consumption of juvenile Nile tilapia exposed to high (sub‐lethal) and low levels of total water ammonia (TAmm). The first series of experiments aimed to determine the effects of high TAmm toxicity on indicators of metabolic rate and whole‐body growth. Results of non‐
acclimation exposures to ammonia suggest that high levels of TAmm (1000, 2000 and 4000 μM) negatively affect oxygen consumption and ventilation rates, with reduced respiratory efficiency at 4000 μM. This effect on oxygen consumption was not present after a 48hr acclimation period to TAmm concentrations. Tilapia grown under the TAmm treatment conditions had significantly reduced weight and length after 84 days at concentrations of 2000 and 4000 μM. The second series of experiments investigated metabolic rate and growth under conditions of low‐level TAmm (75, 150, 300, 600, 1200 and 2400 μM) to determine potential positive effects on growth. The results of these experiments indicated that oxygen consumption was reduced in non‐acclimated fish at concentrations of 75, 150 and 300 μM, which were therefore examined in subsequent growth experiments. This oxygen consumption reduction was not present after 48hrs of ammonia acclimation. ii Tilapia grown at low TAmm (≤300 μM) did not exhibit significant differences in weight, length, condition factor or specific growth rate within the 56‐day experiment. This study demonstrates that high levels of TAmm significantly impair tilapia whole‐body growth. Furthermore, low levels of TAmm (≤300 μM) do not appear to affect growth. In both series, an initial reduction in metabolic rate was noted in non‐
acclimated fish, but was not present after 48hr TAmm acclimation. While fish “recovered” from initial effects of high TAmm on oxygen consumption and ventilation, significant negative effects on growth were noted. This study suggests that tilapia adapt to the initial effects of TAmm through a process that, at high levels, is energetically costly and compromises growth. iii ACKNOWLEDGMENTS To begin, I would like to extend a sincere thank you to my supervisor, Dr. Bruce Tufts. He has provided me with an immense opportunity for higher learning and growth in academia in taking me on as a Masters student in his lab, for which I will be ever grateful. To my committee members, Dr. Yuxiang Wang and Dr. Shelley Arnott, I am grateful for your input and guidance during this process, thank you for your keen encouragement throughout. I would also like to extend my thanks to those members of the lab who have made my time at Queen’s truly enjoyable; thank you Matt, Tom, Joyce, Melanie, Caleb, Jeff and Rush. I would particularly like to thank Cory for his patience with my questions and his dedication to our science, his optimism it is truly inspiring. Thank you also to all my friends at Queen’s who have kept me engaged inside and outside of the lab. There aren’t enough words to express how you’ve helped me develop as a student and as a person. Thank you especially to Alyson; you’ve been my support through many hard times and with me for the best of them over the past two years. Lastly, I cannot begin to express how much the support of my family means to me. I am so truly lucky to have your unwavering support in all my endeavours, through tough times and otherwise. It is to you I dedicate this thesis. Thank you. iv TABLE OF CONTENTS ABSTRACT...................................................................................................................... ii ACKNOWLEDGEMENTS..................................................................................................iv TABLE OF CONTENTS.................................................................................................... v LIST OF FIGURES............................................................................................................ vi LIST OF APPENDICES..................................................................................................... viii LIST OF ABBREVIATIONS............................................................................................... ix CHAPTER 1: INTRODUCTION AND LITERATURE REVIEW...............................................1 CHAPTER 2: MATERIALS AND METHODS..................................................................... 15 CHAPTER 3: RESULTS..................................................................................................... 23 CHAPTER 4: DISCUSSION............................................................................................... 44 REFERENCES.................................................................................................................. 55 APPENDICES.................................................................................................................. 66 v LIST OF FIGURES FIGURE 1: Oxygen consumption rates of juvenile Nile tilapia acclimated for 0 and 48hr in four sub‐lethal total water ammonia treatments..............................................24 FIGURE 2: Ventilation rates of juvenile Nile tilapia acclimated for 0 and 48hr in four sub‐lethal total water ammonia treatments................................................................. 25 FIGURE 3: Mean weight of juvenile Nile tilapia reared over 84 days in four sub‐lethal total water ammonia treatments.................................................................................. 27 FIGURE 4: Mean length of juvenile Nile tilapia reared over 84 days in four sub‐lethal total water ammonia treatments.................................................................................. 28 FIGURE 5: Mean condition factor of juvenile Nile tilapia reared over 84 days in four sub‐lethal total water ammonia treatments................................................................. 30 FIGURE 6: Oxygen consumption rates of juvenile Nile tilapia in low‐level total water ammonia treatment exposure groups........................................................................... 31 FIGURE 7: Oxygen consumption rates of juvenile Nile tilapia in 0 and 48hr in treatment exposure groups........................................................................................... 32 FIGURE 8: Ventilation rates of juvenile Nile tilapia in low‐level total water ammonia treatment exposure groups........................................................................................... 34 FIGURE 9: Ventilation rates of juvenile Nile tilapia in 0 and 48hr in treatment 35 vi exposure groups............................................................................................................
FIGURE 10: Weight of juvenile Nile tilapia reared over 56 days in four different low‐ level water ammonia treatments.................................................................................. 36 FIGURE 11: Length of juvenile Nile tilapia reared over 56 days in four different low‐ level water ammonia treatments.................................................................................. 37 FIGURE 12: Condition factors of juvenile Nile tilapia reared over 56 days in four different low‐level water ammonia treatments............................................................ 39 FIGURE 13: Specific growth rates of juvenile Nile tilapia reared in four different low‐ level water ammonia treatments over 56 days............................................................. 40 FIGURE 14: Oxygen consumption rates of juvenile Nile tilapia reared in four different low‐level water ammonia treatments over 56 days....................................... 41 FIGURE 15: Ventilation rates of juvenile Nile tilapia reared in four different low‐level water ammonia treatments over 56 days......................................................................43 vii LIST OF APPENDICES APPENDIX 1: Growth distribution of series 1 fish.......................................................... 63 APPENDIX 2: Statistical values corresponding to Chapter 3: Results............................ 64 APPENDIX 3: Coughing observations in high sub‐lethal ammonia preliminary experiments................................................................................................................... 65 APPENDIX 4: PIT‐tag surgery protocol for juvenile tilapia < 10g................................... 66 viii LIST OF ABBREVIATIONS ATP..................................................................................Adenosine triphosphate ANOVA...................................................................................Analysis of variance B................................................................................................................Biomass Δ.....................................................................................................Change in unit CF.................................................................................................Condition factor DO..............................................................................................Dissolved oxygen HSD......................................................................Honestly significant differences L...................................................................................................................Length l.....................................................................................................................Litres MOC....................................................................Metabolic oxygen consumption MO2 ..............................................................................Oxygen consumption rate PNH3...........................................................................Partial pressure of ammonia PO2................................................................................Partial pressure of oxygen PIT.........................................................................Passive integrated transponder αO2.........................................................................Solubility constant for oxygen SGR........................................................................................Specific growth rate SEM............................................................................Standard error of the mean T.....................................................................................................................Time TAmm....................................................................................Total water ammonia ix TCA...................................................................................Tricarboxylic acid cycle V.................................................................................................................Volume W............................................................................................................... Weight x Chapter 1: INTRODUCTION AND LITERATURE REVIEW Historically, natural fish populations have been viewed as a limitless resource. Today, fish constitutes the primary source of animal protein for over a billion people worldwide (FAO 2000), providing 16% of the world’s high‐quality consumed protein and contributing a vital food source for human populations (Tidwell and Allan 2001). The demand for fish products continues to increase. Human population growth has exacerbated our dependence on fish harvest, driving investment in fisheries technology and increasing fishing effort. Improvements in global fishing technologies coupled with poor marine fisheries population management are depleting natural marine stocks (Mullon et al. 2005). An alarming portion of wild fisheries are overfished, overharvested or threatened, leading the FAO to declare 52% of marine fisheries to be at their maximum sustainable limits (FAO 2005), with 70% in need of urgent management (MacLennan 1995). The practice of fishing down marine food webs is also contributing to the collapse of marine fisheries (Pauly et al. 1998). The prospect of fisheries losses on this scale raises serious concerns about the implications of future exploitation (Jenkins 2003; Mullon et al. 2005) and presents a bleak outlook for dependant human populations. Aquaculture Growth: Aquaculture, the farming of fish for human consumption or other uses, has been rapidly expanding in response to this crisis. According to the FAO (2006), aquaculture 1 has grown into a multi‐billion dollar industry. Rapid growth in the aquaculture industry has helped to alleviate some of the human dependence on depleted natural fish stocks. The origins of fish culture extend back hundreds of years to small inland pools of fish in Asia, Egypt and central Europe (Pillay and Kutty 2005). Carp and catfish were some of the first fish to be cultured. Carp were reared in the open ponds of China and catfish culture originated in Cambodia. Traditionally, small‐scale fish farming was adopted in rural communities, as a means of human sustenance and also as a component of agricultural feeds. This century has seen unprecedented growth in aquaculture. Production has expanded extensively from Asia and the Middle East to Western nations. The widespread introduction of aquaculture to North American markets began in the 1960s. Production and value of cultured fish doubled in the 1980s (Naylor et al. 2000). Worldwide, fish production and trade is currently estimated at $89 billion USD per annum in exports, employing over 43.5 million people (FAO 2008). Aquaculture is presently the fastest growing food industry, with a global growth rate of 11% per year. Fish culture is also being rapidly adopted in third world countries, growing approximately six times faster in developing nations than in the first world. The growth of aquaculture is contributing to food security and increased nutritional standards (Tidwell and Allan 2001). 2 Challenges for aquaculture production: Increased commercial demand for fish has stimulated research to optimize cultured fish production. Modern commercial aquaculture typically involves large numbers of fish in open water cage culture or closed water systems. Such production is limited by several parameters, including dissolved oxygen, feed costs and water management (Naylor et al. 2000; Wurts 2000; Liao and Mayo 1974). These issues must be refined to address environmental concerns, commercial sustainability, and to optimize fish growth (Crab et al. 2007; Read and Fernandes 2003, Beouf and Payan 2001). Perhaps the greatest problem that must be addressed in modern aquaculture is nitrogenous waste management (Eddy and Williams 1987; Liao and Mayo 1974). Waste products in areas of intense aquaculture production can impact the surrounding environment. These effects can be severe in shallow or confined water bodies (Wik et al. 2008; Naylor et al. 2000). This presents problems in terms of potential damage to the environment, and contributes to negative perceptions of aquaculture from members of the scientific community and general public. Open‐caged facilities based in coastline regions have come under intense scrutiny. Production in these facilities consists of high‐densities of cultured fish or shellfish, which are often fed high levels of antibiotics to combat the spread of disease. These antibiotics subsequently enter the surrounding areas, where they are exposed to plants and animals, which can lead to the selection of antibiotic‐resistant strains. The 3 open nature of these environments also allows fish feed and effluent to accumulate at the base of culture cages, resulting in the eutrophication of local waters (Landesman 1994) and compromised water quality (Eng et al. 1989). In contrast, closed water systems provide an alternative means of production that reduces environmental impacts and allows for greater control of factors that might affect growth. This approach to intensive culture has many benefits, including the reduction of the number of escapees into natural environments, effectively containing fish populations. Closed systems also allow for the direct treatment of wastes, the ability to limit the release of antibiotics to the external environment, and control over water pH, temperature and chemistry. Such fine manipulation of the culture environment helps optimize fish growth. Nitrogenous aquaculture wastes: Nitrogenous waste products can present significant obstacles to fish growth. The rapid accumulation of ammonia (NH3), and to a lesser degree nitrite (NO2‐), can lead to mass mortality. Although nitrite is not usually a major factor in the natural environment, it is a major concern in closed (i.e., recirculating) systems because relatively small concentrations can be lethal (Cheng et al. 2004; Eddy et al. 1983). Nitrite is a nitrogenous intermediate formed during the nitrification of ammonia by bacteria or through the denitrification of nitrate. It acts by disrupting the normal structure of blood haemoglobin, converting haeme groups into nitrite‐bound forms. This results in methaemoglobinemia, a reduction in the oxygen carrying capacity of haemoglobin, 4 which has severe physiological effects (Jensen 1990; Margiocco et al. 1983). Under high‐
density culture conditions where dissolved oxygen concentrations can be compromised, reduced oxygen binding capacity can lead to fish suffocation and mass mortality (Jensen 2003). Ammonia is the major product of teleost excretion (Cheng et al. 2004; Tomasso 1994) and comprises the majority of nitrogenous waste in intensive aquaculture production. It is also the nitrogenous end‐product of amino acid and protein oxidation (Smith and Rumsey, 1976). In fish, ammonia is produced predominantly in the liver through the transamination of amino acids, but it may also be produced in the kidneys, gills, and skeletal muscle (Randall and Wright 1987). Approximately 80‐90% of the nitrogenous waste is excreted by the gills (Goldstein and Forster 1961). In solution, ammonia exists in equilibrium between two forms: the readily soluble ammonia gas (NH3) and the ammonium ion (NH4+), together known as total ammonia (TAmm). Under normal body conditions TAmm will exist mainly in ionic form (>95%), but under conditions of increased temperature or pH, this equilibrium shifts towards formation of NH3 (Wood 1993). The non‐ionic ammonia (NH3) molecule is highly toxic, rapidly permeating gill and tissue membranes and impeding central nervous system function (For review see Evans et al. 2005 and Randall and Tsui 2002). Such disruptions in fish may result convulsions, loss of physical equilibrium, or mortality (Randall and Tsui 2002; Felipo and Butterworth 2002; McKenzie et al. 1993). Ammonia concentrations are regulated by 5 passive diffusion down the partial pressure gradient of NH3 across the gill (Wilson et al. 1994). This gradient is maintained by the simultaneous active excretion of protons from the gills. At the gill surface, protons bind to ammonia molecules, resulting in the formation of ammonium ions. Ammonium (NH4+) is regarded as the less toxic form of ammonia. Due to its larger size and charge, ammonium is much less diffusive and is restricted to movement through water‐filled channels. Excretion of ammonium is coupled to the movement of other ions via H+/NH4+ and Na+/NH4+ exchangers, or to a lesser degree via electrochemical diffusion from the gills (Randall et al. 1995; Wright and Wood 1985). These ions are impermeable to fish biological membranes and remain in the external environment. Thus, the conversion of ammonia (NH3) to ammonium (NH4+) effectively “traps” ammonia outside the fish (Wood 1993). This process removes the toxic form of ammonia from the fish and preserves the critical gradient for NH3 diffusion. Rates of ammonia excretion in fish vary with changes in the catabolism of dietary or structural proteins. The greatest change in total ammonia efflux is seen with changes in protein consumption. When fed balanced diets, fish allocate the majority of their consumed nitrogen to growth and energy production (Wood 1993). If fish consume amino acids in excess of what is required for maintenance and growth, the surplus is either deaminated and then oxidized via the Krebs cycle, or converted to fat and carbohydrate (Wood 1993). Under conditions of starvation, fish break down muscle protein as a substitute and follow a similar process of deamination. Blood ammonia 6 concentrations in fish are affected by a variety of exogenous and endogenous factors. For example, elevated temperatures increase the deamination rate, raising plasma ammonia levels. Similarly, increases in exercise lead to excesses of ammonia production over excretion, elevating blood ammonia (Wood 1998; Wood 1993). Whenever production of ammonia from amino acid deamination and adenylate deamination surpasses the excretory rate, it results in elevated levels of blood ammonia. Excess levels of total ammonia present a major obstacle to intensive fish culture, as high volumes of uneaten feed and feces lead to the accumulation of nitrogenous waste (Tovar et al. 2000; Clark et al. 1984). Unmanaged total ammonia (TAmm: NH3 + NH4+) concentrations in aquaculture are known to compromise fish health, retard growth and cause mortality (Person‐Le Ruyet et al. 1997; Meade 1985; Balarin and Haller 1982; Smart 1976). In flow through systems, where ample fresh water and oxygen are supplied to clear nitrogenous compounds, ammonia does not present a significant obstacle to fish culture. In contrast, closed, recirculating culture systems can rapidly accumulate nitrogenous waste compounds and require nitrifying biofilters. These biofilters consist of bacteria which are fixed to media and grow in the presence of inorganic nitrogenous compounds (Lyssenko and Wheaton 2006). The bacteria utilize ammonia and nitrite in their metabolic processes, removing them from the water. To date, research into the effects of ammonia on cultured fishes has primarily focused on its detrimental effects as a waste product (Eddy 2005; Randall and Tsui 2002; Arillo et al. 1981). Consequently, “ammonia regulation” in aquaculture is widely 7 approached with the objective of complete ammonia removal. It is also important, however, to consider the effects of ammonia at levels below toxic concentrations. Potential benefits of nitrogenous wastes for fish growth: Paradoxically, Linton et al. (1997) noted that exposure to low levels of ammonia stimulated growth in juvenile rainbow trout without an increase in food consumption. Fish in Linton’s study were grown in conditions of elevated ammonia (70µM TAmm) at natural water temperatures, and gained significantly greater weight than ambient water controls. Furthermore, ammonia exposed fish had an increased nitrogen and energy conversion efficiency with higher nitrogen retention at a lower metabolic cost (Linton et al. 1997). Linton observed that fish exposed to elevated ambient water ammonia were retaining greater overall levels of nitrogen, in the form of protein, and utilizing it to enhance growth. The authors proposed that sub‐lethal levels of water ammonia elevated ammonia concentrations in fish plasma, triggering ammonia detoxification mechanisms, which in turn stimulated protein turnover rates and lead to a higher retention of nitrogen. Linton hypothesized that this increased retention of nitrogen resulted from the incorporation of ammonia into amino acids such as glutamine. This was reflected in the results by elevated levels of glutamine in the livers of ammonia‐
exposed fish when compared to the control group (Linton et al. 1997). 8 In a subsequent study, Linton observed that trout exposed to ammonia over the course of 14 months retained up to 10% more nitrogen than control fish (Linton et al. 1998), though this was not statistically significant. Linton suggested that this stimulatory effect of ammonia exists while fish are not subjected to excessive temperature or toxicological stress (Linton et al. 1998). This may provide a shift in energy allocation whereby fish down‐regulate metabolism by limiting protein catabolism and allocating more energy to ammonia detoxification via amino acid formation (Linton et al. 1998). Recent studies have noted beneficial effects of low levels of total ambient ammonia (Madison et al. 2009; Wood 2004, Ip et al. 2001) and its potential to enhance fish growth. Wood (2004) grew juvenile rainbow trout at fixed temperatures of 15 and 6.5 °C and at concentrations of 0, 70, and 220µM (TAmm). He observed significant increases (compared to controls) in mass, food conversion and protein content after 70 days at 70µM (TAmm) in fish grown at 15°C. However, at 6.5°C no significant effect on growth was observed at 70µM (TAmm). Instead, significant increases in growth and protein accretion were noted at 220µM (TAmm). Wood highlighted two possible explanations. First, the effect of ammonia on growth may be a product of the relative concentration of NH3, which increases with temperature relative to NH4+. Second, conflicting results upon examining the significance of growth at days 42 and 71 at 6.5°C suggested that the growth effects may only occur after long term ammonia exposure. Wood also suggested that increased protein accretion without an alteration in food 9 consumption may be the result of reduction of metabolic costs from ammonia exposure. Madison et al. (2009) observed significant reductions in the metabolic rate of juvenile walleye when exposed to concentrations of 75 and 150µM TAmm. These findings appear to conform to previous studies noting reductions in metabolic activity, and are in agreement with hypotheses put forward by Beaumont et al. (1995). Beaumont et al. (1995) suggested that exposure to ammonia reduces the routine metabolic rate through the interference of aerobic metabolism by slowing or impairing oxidative phosphorylation of within the TCA cycle. This would lead to a shift toward anaerobic metabolism, whereby glucose and glycogen would be utilized alternatively through glycolysis. This strategy allows for improved ammonia detoxification and leads to a storage of nitrogenous by‐products (Randall and Tsui 2002). The above hypothesis was further supported by the findings of Madison et al. (2009), which included a significant depletion of whole‐body glycogen and tissue glucose levels with ammonia exposure. Madison et al. (2009) further proposed that the reduction in metabolic rate seen in fish at 75 and 150µM TAmm may have been the result of a threshold “calming effect” of plasma ammonia, while concentrations at 300µM TAmm induced stress. This effect of reduced metabolic rate is hypothesized to be the result of nitrogenous detoxification through the amination of glutamate by glutamine synthetase to form glutamine (Felipo and Butterworth 2002; Randall and Tsui 2002). Over‐activity of this detoxification mechanism exhausts stores of brain glutamate and 10 ATP, impairing neurotransmission and driving the synthesis of glutamine (Monfort et al. 2002). In moderation, this process may calm fish by reducing the release of compounds triggered by physiological stress (Madison et al. 2009). Juvenile walleye grown over 56 days at 18°C in 75 and 150μM TAmm demonstrated significantly higher levels of whole‐body protein and a significantly higher growth rate during the 14‐42 day period, as compared to controls (Madison et al. 2009). Elevated rates of protein turnover were also observed at these concentrations. Increases in fish internal ammonia activate detoxification mechanisms. In ammoniotelic fish, these mechanisms may enhance amino acid synthesis and reduce protein catabolism (Wood 2004; Ip et al. 2001; Mommsen and Walsh 1992). Increases in internal ammonia may result from several environmental factors, including low water pH and high levels of ambient water ammonia, and lead to a reduced gill transepithelial ammonia gradient (Beaumont et al. 2000). Under normal growth conditions, fish maintain stable levels of protein (as a percentage of wet weight). However, Linton et al. (1997) and Wood (2004) have hypothesized that marginally‐increased concentrations of ammonia in fish plasma result in modifications of the detoxification pathway, leading to increased protein accretion with the incorporation of nitrogen. Such results highlight the potential for ammonia to alter fish metabolism. This could lead to protein accretion at critical levels of TAmm in early life stages, enhancing the growth of cultured fish. Combined, these studies challenge the conventional approach to ammonia removal and 11 suggest that regulated low ammonia concentrations may actually benefit fish growth in aquaculture. Tilapia aquaculture: Due to their attributes as a fish species with a wide range of tolerances to environmental conditions, rapid growth rate, and wide range of diet options, tilapia (family Chichlidae) have emerged as an important species in aquaculture (El‐Sayed 2006; Popma and Lovshin 1995). Tilapia have been successfully reared in fresh, brackish, and salt water environments and are able to subsist on plant and omnivorous diets. High density populations often present challenges in terms of maintaining high water quality; however, tilapia have proven to be remarkably robust under compromised conditions. Tilapia ammonia tolerance has been documented to be as high as 2.4mg l‐1 (LC50, 48h) in unacclimated fish, and 3.4mg l‐1 (LC50, 48h) in fish acclimated to a sub‐lethal level of ammonia (Redner and Stickney 1979). Remarkably, dissolved oxygen tolerances of tilapia have been recorded to as low as 0.1mg l‐1 (Stickney 1986), considerably lower than other cultured fish. Furthermore, while sensitive to temperatures below 10°C, tilapia may be tolerant of temperatures up to 42°C (Mires 1995), with an optimal growth range of 22 to 29°C. This variety of tolerances has made it an ideal fish for growth in rural regions. Tilapia have been identified as one of the first fish cultured in ponds in Egypt (Pillay and Kutty 2005). Originally from Africa, tilapia species have spread to countries all around the world with particular success in tropical and sub‐tropical regions. Though 12 many species of tilapia exist, aquaculture is generally dominated by only a few species. Of these, the organized culture systems of Mozambique tilapia (Oreochromis mossambius) appear to have originated in African regions and spread to Asia in the 1940s. This species has since been surpassed by the aquaculture of Nile tilapia (Oreochromis niloticus), which surged in the 1960s, and currently make up the majority of aquaculture production worldwide (Edwards et al. 2000). Tilapia are of immense socio‐economic value, with a total international production of over 2,000,000 metric tons in 2004 (Fitzsimmons 2006). Nile tilapia comprises 80% of total production. Tilapia’s tolerance for poor water and habitat quality means they are appropriate for culture in developing countries (El‐Sayed 1999). With their omnivorous diet, tilapia also represent an appropriate option to minimize the impact of aquaculture feed production on wild fish stocks (i.e., fishing down the food web) (Pauly and MacLean 2003; Naylor et al. 2000; El‐sayed 1999). Current trends towards intensive commercial farming of fish, coupled with enhanced marketing, have created an increased demand for tilapia. Tilapia production has increased by 390% from 1990 to 2002, and is currently increasing in production at 12.2% annually (El‐Sayed 2006). In view of the previous discussion, it is clear that ammonia can have an important impact on fish in aquaculture situations. To date, however, there is only limited information about the impacts of ammonia on tilapia. 13 Thesis objectives:
This thesis examines the effects of ammonia on the growth of tilapia under aquaculture conditions. The first series of experiments in this study addressed the effect of sub‐lethal levels of TAmm on juvenile Nile tilapia growth and oxygen consumption under high‐
density culture conditions. The effects of a broad range of sub‐lethal concentrations of ammonia will be examined. The second series examined lower levels of ammonia that have previously elicited positive growth effects in rainbow trout (Oncorhynchus mykiss) and Walleye (Sander vitreus). Nile tilapia will be grown from a juvenile stage at these levels of TAmm, under similar intensive culture conditions as in series 1, to determine whether low levels of TAmm have a positive impact on growth. Based on the results of previous studies involving other species, it was hypothesised that high levels of TAmm would be detrimental to the growth of juvenile Nile tilapia in series 1. For series 2, it was hypothesised that positive effects of low levels of TAmm seen in other species may also be observed in Nile tilapia. Together, these experiments generate practical insight into the management of TAmm in tilapia culture by assessing the effects of TAmm on metabolism and growth. This thesis attempted to establish levels of TAmm that may optimize the aquacultural production of Nile tilapia, arguably the most important emerging aquaculture species today. 14 Chapter 2: MATERIALS AND METHODS Two experimental series were performed to examine the effect of sub‐lethal levels of ammonia on juvenile Nile tilapia metabolism and growth. Experimental Animals: Juvenile Nile tilapia (Oreochromis niloticus, each approximately 5g) were acquired from North American Tilapia (Elmira, ON, Canada), a regional aquaculture facility. Fish were transported to the Biology Department Animal Care facility at Queen’s University (Kingston, ON, Canada) and transferred to a 1000 litre flow‐through holding tank supplied with de‐chlorinated City of Kingston tap water. Holding tank conditions were maintained at 26±1°C, pH 7.6 with a dissolved oxygen concentration of > 6.0 mg l‐1. Fish were held on a 14h:10h light:dark photoperiod and were fed a diet of commercial feed pellets (Corey Feed Mills Ltd, NB). Fish were fed to satiation once daily and were held in these conditions for approximately 50 days prior to experiments. Series 1 The first series examined the effects of high‐levels of ammonia on oxygen consumption, ventilation rate and growth. The purpose of this series was to determine the upper threshold of ammonia tolerance in Nile tilapia, and to examine the physiological effects of ammonia. 15 Toxicity range‐finding experiment: A preliminary toxicity range‐finding experiment was undertaken to establish the effective total ammonia (TAmm) concentration eliciting a physiological response. Groups of 4 fish (each approximately 5g) were introduced to aerated chambers at 25±1°C at different concentrations of ammonia delivered by adding NH4Cl to fresh treatment tank water and stirring until dissolved. The effective [TAmm] was characterized by the concentration that led to a loss of orientation, “coughing”, surface gulping, or erratic swimming inside test chambers after 48hr exposure. The upper limit to toxicity was observed at 8 mM TAmm. In subsequent experiments fish were introduced to chambers at lower concentrations of TAmm. These exposure concentrations were reduced from the observed limit in multiples of 1000 μM until no behavioural response was present. The highest concentration of ammonia which did not elicit an acute (i.e., immediate) behavioural response was used in the high‐ammonia growth and oxygen consumption experiments of series 1. Oxygen consumption experiment: Two sets of experiments were conducted to examine the impacts of high levels of ammonia (as determined by the range finding experiment) on metabolism. Experimental fish were divided into two groups, non‐acclimated and acclimated. In the first set, individual fish were added without any acclimation period to 4L jar respirometers (Cech 1990) with pre‐mixed concentrations of 0, 1000, 2000, 4000 μM 16 TAmm in water. Fish in the first treatment group were transferred directly from the flow‐
through (0‐12 μM TAmm) holding tank to a respirometry chamber for individual experimentation (Total N = 32 fish). This non‐acclimated group was used to determine the acute effects of the ammonia treatments on fish ventilation rate and oxygen consumption. Fish in the second group were introduced to a 75L aquarium at their respective treatment concentrations for 48hrs prior to experimentation in the respirometry chambers (N = 32). Acclimation aquaria were supplied with aerated, mechanically filtered water in recirculating tanks, with the same conditions as the holding tank. Fish were subsequently transferred to respirometry chambers for experimentation. This acclimated group was used to observe the non‐acute effects of the ammonia treatment. In all experiments, respirometry chambers were filled with fresh aerated water; measured NH4Cl was stirred into solution. The chambers were then sealed upon the introduction of fish until the end of the experiment. Dissolved oxygen readings were recorded at 0 and 120 minutes. Oxygen consumption was measured using a dissolved oxygen combination probe and meter (Yellow Springs Instruments, Model 55, Yellow Springs, OH, USA). Respirometry jars followed the design of Cech (1990) and were maintained in water at 26.5±0.5°C to control for temperature effects (ie minimize temperature fluctuations). Immediately following the trial, fish were measured for weight (Ohaus portable plus scale, to 0.01g) and length (to 0.1cm) for the calculation of oxygen consumption (mg kg‐1 hr‐1) and condition factor. 17 Growth experiment: Experiments to determine the effect of high‐levels of ammonia on juvenile Nile tilapia growth were conducted over a period of 84 days beginning in mid‐March 2008. First, 80 fish from the holding tank were randomly divided into one of the four 75 litre treatment tanks (0, 1000, 2000 and 4000 μM TAmm). Prior to the experiment, fish were individually weighed (Ohaus portable plus scale, to 0.01g) and total length was measured (to 0.1cm) to establish day 0 parameters. The same variables were then measured on a weekly basis until the conclusion of the experiment. The growth treatment tanks were each supplied with airstones, heaters (Ebo‐
Jager Model TS 125W), recirculating pumps and filters (AquaClear 200). Each tank was supplied with de‐chlorinated Kingston tap water maintained at 26±1°C. Water was changed regularly to maintain clarity and treatment concentrations. Treatments consisted of four groups: [TAmm] = 0, 1000, 2000, 4000 μM TAmm. Treatment concentrations were obtained by adding NH4Cl to fresh treatment tank water and stirring until dissolved. Tank pH was monitored at least once a week with dissolved oxygen and temperature monitored daily to ensure minimal fluctuation. Series 2 A second experimental series, following the same approach as the first series, was conducted to observe the effects of low levels of ammonia on metabolic rate and 18 growth. More specifically, this series was conducted to determine if low‐levels of ammonia enhanced growth through a reduction in metabolism. Oxygen consumption experiment: The respirometry experiments undertaken in series 2 examined the effects of ammonia on metabolic rate and followed an identical experimental design as in series 1, with one exception: the range of low‐levels of ammonia was as follows 0, 75, 150, 300, 600, 1200 and 2400 μM TAmm. The experiments were again divided into non‐acclimated (Total N = 56 fish) and acclimated groups (Total N = 32 fish). The full low‐ammonia range (0‐2400 μM TAmm) was used for the non‐acclimated group to determine the effects of the ammonia treatments on fish ventilation rate and oxygen consumption. In subsequent experiments with acclimated fish, treatment groups were limited to levels of ammonia that demonstrated an acute effect on metabolism (0‐
300 μM TAmm) in previous (non‐acclimated) experiments. The methodology for the respirometry aspect of these experiments followed a similar format as in series 1. In these experiments, however, oxygen consumption was measured using a water‐jacketed Radiometer oxygen electrode (Radiometer E5046, Denmark) maintained at 26.5°C. Concentrations of ammonia that elicited an effect on metabolism were utilized in the subsequent series 2 growth experiment. 19 Growth experiment: The low‐ammonia growth experiment followed a similar design to series 1. The experiment was conducted over a period of 56 days beginning in mid‐August 2008 to examine the impact of various low‐levels of ammonia on Nile tilapia growth. A total of 160 fish from the holding tank were randomly divided into one of the eight 75 litre treatment tanks ([TAmm] = 0, 75, 150, 300 μM, in duplicate). Fish were measured on a bi‐
weekly basis until the conclusion of the experiment (weeks 0, 2, 4, 6 and 8). Each fish was uniquely identified by passive integrated transponder (PIT) tags, surgically inserted into the body cavity under anaesthesia prior to the beginning of experimentation. The 12mm PIT tags (TX1441SST, 134.2khz) and reader system (Biomark, Pocket reader EX, Biomark, Idaho, USA) were employed to generate bi‐weekly data for growth analysis from measured lengths and weights. The growth treatment tanks were run in duplicate and set‐up in a manner identical to series 1. The water was changed daily to maintain clarity and treatment concentrations. Treatment concentrations were obtained by adding dissolved NH4Cl solution to fresh treatment tank water via Mariotte bottle. Tank pH was monitored at least once a week. Dissolved oxygen and temperature were monitored daily to ensure minimal fluctuation. 20 Analytical techniques: In both series 1 and 2 water pH was read using a pH meter (Fisher Scientific, Accumet Basic AB15 pH meter). Total water ammonia (NH3 and NH4+) was quantified based on the indophenols blue method of Ivancic and Degobbis (1984). Calculations and statistical analysis Respirometry measurements: Oxygen consumption rates (MO2) of both series were calculated from the rate of change of PO2 (ΔPO2/t, torr h‐1) values over the duration of experiments: MO2 = [(ΔPO2/t)αO2V]/B, (1) where αO2 (µmol l‐1 torr‐1) is the solubility constant for O2 in water for the experimental conditions (Boutilier et al. 1984), V is the volume of respirometer, t is time and B is the total biomass of a fish. Ventilation rates were monitored so as to not cause behavioural disturbances (i.e.,, observations were made at a distance and under stable light conditions with no abrupt movement in the visual field of the fish) while fish remained in the respirometry equipment. Ventilation rate was defined as the number of complete brachiostegal membrane movements per minute. 21 Growth measurements: Specific growth rate (SGR, % wet weight gain day‐1) over an experimental interval (t, days) was calculated from individual fish wet weight (Wn, g): SGR = [(lnWn+1‐lnWn)/t]100 (2) Condition factor (CF) was calculated from individual weight (W) and length (L, cm) as per Busacker et al. (1990): CF = (W/L3)100 (3) Statistical analysis: Data are displayed as means ±SEM (N) where N = the number of fish samples. Series 1 oxygen consumption, ventilation rate, weight, length, condition factor and specific growth rate data were analyzed by using a two‐way analysis of variance (ANOVA) to determine whether there were significant differences from Day 0, as well as among experimental treatment groups. Series 2 oxygen consumption, ventilation rate, weight, length, condition factor and specific growth rate data were analyzed using a repeated measures two‐way analysis of variance (ANOVA). Differences in variables in each series were detected using a Tukey‐Kramer HSD post hoc test. All statistical testing was performed using JMP 7 (SAS Institute), and the level of significance (α) for all tests was 0.05.
22 Chapter 3: RESULTS Series 1 Oxygen consumption experiment: In non‐acclimated fish, oxygen consumption (mg kg‐1 hr‐1) was significantly lower at 2000 and 4000μM total ammonia (TAmm) relative to the control group (Figure 1). For fish acclimated to different ammonia levels for 48hrs, no significant difference in MO2 was observed relative to control fish. When comparing between the acclimated and non‐acclimated series, the oxygen consumption rate of acclimated control fish was significantly lower than that of non‐acclimated control fish. The reverse was observed between ammonia‐exposed fish in the two experimental series with the acclimated fish exhibiting higher oxygen consumption rates than their corresponding non‐acclimation treatments. This difference was significant for the 4000µM TAmm treatment. In the non‐acclimated series, ventilation rates (ventilations minute‐1) were significantly lower in the 2000μM TAmm treatment group compared to the control group (Figure 2). Ventilation rates at 4000μM TAmm were significantly greater than at 2000μM TAmm, but not significantly different from 1000μM TAmm or control fish. In the acclimated series, there were no significant differences in ventilation rates among treatment groups. When comparing acclimated and non‐acclimated experimental series, significant differences were only observed at 2000μM TAmm, with lower ventilation rates in non‐acclimated fish. 23 24 25 Growth experiment: Fish weight increased throughout the duration of the experiment for all treatment concentrations. By 42 days, the control, 1000, and 2000µM TAmm groups had attained a significantly greater weight than the pre‐experimental control. By 56 days all treatment groups were significantly greater in weight than the pre‐experimental control. The 4000µM TAmm treatment weight at day 56 was significantly less than the 1000µM TAmm treatment, and was significantly lower than all treatments at day 84 (Figure 3). Fish length in the 4000μM TAmm treatment was significantly less than the control group (Figure 4). At 28 days, the control group fish were significantly longer than the pre‐experimental control fish. At 42 days all treatment groups had attained significantly greater length than the pre‐experimental control. Lengths of the 4000μM TAmm treatment were significantly smaller than the control and 1000μM TAmm treatments from 56 days to the conclusion of the experiment. 26 27 28 At day 42, the 4000μM TAmm treatment group was observed to have a significantly lower condition factor than the pre‐experimental control (Figure 5). However, no further effect of the TAmm treatments on condition factor was observed. Series 2 Oxygen consumption experiment: In the non‐acclimated experimental series, oxygen consumption rates were significantly lower in 75, 150, and 300µM TAmm treatments relative to the control group (Figure 6). However, oxygen consumption at higher concentrations (600, 1200 and 2400μM TAmm) was not significantly different from the control. Total ammonia concentrations yielding significant reductions in oxygen consumption were then run in a second series with a 48hr period of acclimation to treatment concentrations. For 48hr acclimated fish, no significant differences were observed between the ammonia treatment groups and the control group (Figure 7). When comparing between acclimated and non‐acclimated experimental series, all 48hr acclimated treatment groups had lower oxygen consumption rates than their corresponding non‐acclimated groups. The non‐acclimated control exhibited significantly higher oxygen consumption than the acclimated control.
29 30 31 32 Low concentrations of total water ammonia had no effect on ventilation rate in non‐acclimated treatments (Figure 8). However, ventilation rates at 1200 and 2400μM TAmm were significantly reduced relative to control fish and all other treatments. The ventilation rates of 300μM TAmm fish were also significantly lower than those of 75 and 150μM TAmm treatments. In the acclimated series, there were no significant differences in ventilation rates among any of the treatment groups (Figure 9). Oxygen consumption rates for non‐acclimated fish in 75, 150, and 300µM TAmm exposures were significantly lower than those in control fish (Figure 6). These concentrations were used in subsequent growth experiments. Growth experiment: After 42 days, the mean weights in all treatment groups were significantly greater than that of the initial control group. No significant differences between the mean weight of the control group and TAmm treatment groups were observed throughout the 56‐day growth experiment (Figure 10). All treatment groups attained a significantly greater length than the pre‐
experimental control by 42 days (Figure 11) with the exception of the 150μM TAmm treatment, which did not have a significantly greater mean length until day 56. 33 34 35 36 37 No consistent trend was observed of TAmm treatment on condition factor (Figure 12). At day 56 the condition factor of the 300µM TAmm treatment was significantly greater than those of the control and 150μM TAmm treatment groups. However, there was no significant difference between the treatment groups and pre‐experimental control group that persisted throughout the growth period. Overall, no consistent trend of total water ammonia was observed on specific growth rate (Figure 13). However, fish in the 300µM TAmm treatment at the 43‐56 day interval had a significantly reduced specific growth rate compared to the 150μM TAmm treatments. Growth respirometry experiment: At day 14 the oxygen consumption rate of the 300µM TAmm treatment was significantly reduced relative to both the 0µM and pre‐experimental control (Figure 14). At day 28 oxygen consumption in the 300μM TAmm treatment was significantly lower than that of the pre‐experimental control. At day 42 oxygen consumption in the 75 and 150µM TAmm treatments were significantly lower than the pre‐experimental control. All oxygen consumption measurements on day 56 were significantly lower than the pre‐
experimental control. 38 39 40 41 Total water ammonia treatment had no effect on fish ventilation rates compared to the control throughout the experiment (Figure 15). However, the ventilation rates of all treatment groups were significantly lower than that of the pre‐experimental control group throughout the growth period. 42 43 Chapter 4: DISCUSSION This is the first study to examine the effects of sub‐lethal levels of total ammonia on growth and metabolism in aquaculture‐reared Nile tilapia. This is also the first study to thoroughly examine the time course of the effects of ammonia on growth and metabolism in fish. Results from these experiments using juvenile Nile tilapia support previous findings that total water ammonia (TAmm) has a negative effect on fish whole‐
body growth (Lemarié et al. 2004; Rasmussen and Korsgaard 1996; Beamish and Tandler 1990; Robinette 1976). This study also shows that the suppressive effect of ammonia on oxygen consumption in Nile tilapia is limited to a narrow range of concentrations and a limited exposure period. Series 1: High concentrations of sub‐lethal ammonia Oxygen consumption experiment: Results from respirometry experiments indicate that high sub‐lethal levels of TAmm lower resting metabolic rate in non‐acclimated juvenile Nile tilapia. Upon initial exposure to sub‐lethal TAmm concentrations, Nile tilapia exhibited a suppression of oxygen consumption (Figure 1). This may be the result of initial exposure to the toxic environment and the rapid onset of physiological changes. Person‐Le Ruyet et al. (1998) noted that in as few as 15 minutes after exposure, ammonia was present in the blood 44 plasma of turbot and seabream juveniles introduced to increased concentrations of water ammonia. Sub‐lethal levels of ambient water ammonia lead to the disruption of the normal NH3 gradient where ammonia normally passes from the bloodstream to the surrounding waters via passive diffusion across the gills. The reversal of this gradient prevents the diffusive outflow of NH3, leading to an increase in internal ammonia levels. Without regulation, this can lead to toxic levels of total ammonia concentrations in the blood. The lowering of resting metabolic rate upon exposure to ammonia may result from an initial suppression of aerobic metabolism (Beaumont et al. 1995). Beaumont and colleagues (1995) speculate that this may result in a shift towards anaerobic energy production as a mechanism to support routine energy requirements while reducing the production of internal metabolically‐derived ammonia. This mechanism agrees with our findings for the exposure of unacclimated juvenile Nile tilapia. Future studies should include measures of blood lactate throughout the exposure period of Nile tilapia to ammonia in order to confirm that anaerobic metabolism is increased under these conditions. In the present experiments, the 2000μM TAmm treatment lowered the rate of ventilation. However, higher concentrations elicited a clear ventilation stress response in the form of increased ventilation rate (Figure 2) and fish “coughing” (data not shown). Interestingly, while ventilation rate returned to near control levels at the highest concentration, this was not coupled with an increase in oxygen consumption. This may result from a disruption of the aerobic pathway either by the disruption of the TCA cycle 45 in favour of anaerobic metabolism (Beaumont et al. 1995), or through the reduced efficiency of respiratory surfaces. The gills are frequently the first organ to respond to unfavourable environmental conditions. In a previous study, Benli and colleagues (2008) observed that tilapia exposed to elevated ammonia concentrations (2mg l‐1 TAmm) exhibited gill hyperplasia, telangiectasis on lamella and hyperaemia on gill epithelium. Respiratory surfaces that are compromised in this fashion could disrupt routine gas exchange and other gill functions (Miron et al. 2008). Benli and colleges (2008) also noted that tilapia exhibited cloudy swelling and hydropic degenerations of the liver, a critical site for metabolic function. It is interesting to note that in 48hr acclimated fish, the effect on oxygen consumption is no longer apparent (Figure 1). This lack of effect of ammonia on oxygen consumption after 48hr coincides with a reduction or loss of effect on ventilation rate (Figure 2). Together these observations suggest that, after the 48hr acclimation period, the initial effect of total ammonia on the resting metabolic rate in tilapia is no longer present, or has been compensated for. Oxygen consumption rates in ammonia‐
acclimated fish are comparable to those of the controls (0 and 48hrs). However, oxygen consumption was greater after 48 hours of exposure when compared to the unacclimated treatments, suggesting a possible compensatory increase in metabolic rate to meet physiological demands. Person‐Le Ruyet et al. (1998) noted a similar stabilization effect in turbot and seabream juveniles exposed to ammonia. The authors noted that after exposure to elevated concentrations of ammonia, fish plasma TAmm 46 stabilized. This was followed by a return to sub‐ambient levels within 1‐3 hours (Person‐
Le Ruyet et al. 1998). The results of the current experiment provide evidence of Nile tilapia acclimation to elevated ammonia levels in as few as 48hrs. This acclimation is interesting to note in light of work done by Redner and Stickney (1979) who demonstrated that lethal concentrations of ammonia for unacclimated Tilapia aurea could be tolerated by fish previously exposed to moderate concentrations of ammonia for 35 days. In addition, Redner and Stickney (1979) noted that exposure to acute and sub‐lethal concentrations of ammonia resulted in capillary congestion, haemorrhaging and other symptoms of gill abnormalities. The authors suggest this acclimation may be the result of the inhibition or reversal of nitrogen metabolism, a hypothesis also put forth by subsequent studies in trout (Wood 2004; Ip et al. 2001; Mommsen and Walsh 1992). Taken together, these studies provide evidence for mechanisms of acclimation in conditions of high ammonia stress. Results from the current study indicate that this acclimation to elevated ambient ammonia levels may take effect within 48hr in juvenile Nile tilapia. Growth experiments: The results of the initial experiments in this thesis suggest that sub‐lethal levels of TAmm have a negative effect on the growth of juvenile Nile tilapia under aquaculture conditions by suppressing gain in fish weight and length relative to controls (Figures 3 and 4). At the highest ammonia concentration (4000μM TAmm), both fish weight and 47 length were significantly reduced compared to other treatments. A similar negative trend was observed at 2000μM TAmm. No clear difference in fish condition factor was observed among treatments (Figure 5). These results suggest that TAmm does not specifically impair fish weight or length in isolation from one another, but instead impairs overall body growth. These findings may be explained by previous studies demonstrating that ammonia toxicity disrupts the production of energy resources and can result in a reduction of 68% of potential energy output (Zieve 1966). It follows that energy that is normally allocated to optimal growth is shunted towards the detoxification process, with the detoxification of 1mol of ammonia requiring 2mol of ATP (Randall and Tsui 2002). The negative effect of ambient ammonia concentrations on weight gain has been noted in several species, such as seabass (Lemarié et al. 2004), juvenile turbot (Rasmussen and Korsgaard 1996), lake trout (Beamish and Tandler 1990) and channel catfish (Robinette 1976). Our results confirm that sub‐lethal levels of ammonia have similar effects on juvenile Nile tilapia growth, with reduced gains in weight and length.
Series 2: Low concentrations of ammonia Oxygen consumption experiment: Oxygen consumption rates of non‐acclimated fish in 75, 150, and 300µM TAmm exposures were reduced compared to control, demonstrating a reduction in metabolic 48 rate (Figure 6). Interestingly, ventilation rate measurements do not show a significant suppressive effect below 600µM TAmm, but there was a reduction in ventilation above this threshold (Figure 8). This suppression of metabolic rate may be the result of the increased activity of glutamine synthetase, a detoxification mechanism. Glutamine synthetase mediates the amination of glutamate and NH3 in the formation of glutamine. While excess activity though this pathway consumes vital brain ATP and glutamate resources, and can result in neurotransmission failure, the severity of this effect varies with the degree of ammonia exposure (Mommsen and Walsh 1992). Madison et al. (2009) propose that when exposed to moderate levels of ammonia, fish may be “calmed” by this mechanism through reduced release of stress‐induced compounds. Consistent with this hypothesis, our results suggest that Nile tilapia exposed to levels of <300µM TAmm experience reduced oxygen consumption, but above this concentration, the effect is not observed. Such a reduction is consistent with experiments by Madison and colleagues (2009), who noted a similar reduction of metabolic rate in walleye exposed to concentrations above 150µM TAmm during respirometry experiments. Madison and colleagues (2009) observed an increase in specific growth rates in fish grown between 75 and 150µM TAmm, and speculate that juvenile walleye exposed to these levels accumulate sufficient concentrations of plasma ammonia to elicit a threshold calming effect. Madison et al. (2009) further suggest that this growth effect was not seen at higher concentrations (300µM TAmm ) due to overriding physiological stresses resulting from increased plasma ammonia. 49 After 48hrs of exposure, the effect of reduced metabolic rate is no longer present. These results are consistent with our findings above, where the initial toxicological effects of oxygen consumption and ventilation at sub‐lethal concentrations of total ammonia appeared to be less severe or absent after 48hrs (Figures 7 and 9). This again highlights the possibility of a response where aerobic metabolism is compromised upon initial exposure to TAmm, but is restored within a 48hr period. Growth experiment: The results of low‐range TAmm growth experiments suggest that concentrations of 300µM TAmm or below do not affect juvenile Nile tilapia weight or length under aquaculture conditions (Figures 10 and 11). Accordingly, no major effect was seen on fish condition factor or specific growth rate (Figures 12 and 13). Though these concentrations initially resulted in reduced oxygen consumption in respirometry experiments, they did not have a significant effect on growth. Despite ammonia being present in concentrations high enough to elicit a metabolic response, detoxification mechanisms, if active, do not appear to interfere with growth potential in juvenile Nile tilapia at these concentrations. While negative growth effects were not noted at these concentrations, it is also important to acknowledge that there was also no major positive effect from low levels of ambient ammonia. Previous studies on rainbow trout (Wood 2004) and walleye (Madison et al. 2009) growth in similar concentrations of ammonia have demonstrated a potential for enhanced growth, but are not conclusive 50 with regards to the concentration range or duration of exposure necessary to elicit such effects. Wood (2004) conducted two series, each over 71 days. The first, at 15°C, resulted in significant weight gain, as well as increases in condition factor and protein production, at 70µM TAmm. The second series, conducted at 6.5°C, produced similar effects at 225µM TAmm. Wood (2004) concluded that though different temperatures and concentrations existed between the two series, a comparable PNH3 between the experiments may be responsible for the increased protein synthesis and/or a reduction in metabolic costs, resulting in the enhanced growth at these concentrations. Madison and colleagues (2009) observed increased growth rates, higher protein turnover and more whole‐body protein in juvenile walleye exposed to concentrations between 75 and 150µM TAmm at 18°C after 42 days of exposure. Importantly, Madison and colleagues (2009) noted that fish exposed to these low concentrations of ammonia exhibited reduced oxygen consumption. They suggest that these concentrations of water ammonia may reduce metabolic demand, thus augmenting protein synthesis. Alternatively, they suggest that these water ammonia conditions may modify the gill‐
ammonia gradient, leading to a build‐up of internal ammonia available for protein synthesis. Though the concentrations of ammonia in the aforementioned studies demonstrate some benefits to fish growth, they do not appear to have affected the juvenile Nile tilapia in the current study. A study with Nile tilapia grown on a diet of 51 duckweed (El‐Shafai et al. 2004) reported that fish grown in control, 2.5, 5, 7.5 and 10 mg N l‐1 (control, 147, 294, 441, 588 µM TAmm) experienced reduced specific growth rates in concentrations above the ~147 µM TAmm treatment. This result was not corroborated by our study, despite comparable water temperature and pH. Such a discrepancy may be a function of the smaller sample size (N = 4) in the El‐Shafai and colleagues (2004) study compared with those in the current study. Furthermore, it is also possible that the differing feed types between these two studies (duckweed diet vs. high‐performance aquaculture feed) could have contributed to these discrepancies. Growth respirometry experiment: Results from respirometry experiments on juvenile Nile tilapia over the 56‐day growth period further verify a lack of major effect of TAmm on aerobic metabolism (Figures 14 and 15). This result is in agreement with the absence of an effect of low levels of ammonia on whole‐body growth discussed above. However, it is interesting to note that at 300 µM TAmm, significant differences from the pre‐experimental and day 14 controls suggest that if an effect is present at this concentration, it is a transient effect that may warrant further study. This study is the first to evaluate Nile tilapia growth under regulated levels of total water ammonia in aquaculture conditions. These experiments have demonstrated that high (sub‐lethal) levels of total ammonia have a negative effect on juvenile Nile tilapia growth, but it is not yet definitively clear by what mechanism this occurs. The rate of oxygen consumption appears to be altered upon initial exposure of fish to 52 elevated ammonia conditions. However, normal oxygen consumption rates resume soon after, highlighting an adaptive mechanism to initial toxic effects. Although oxygen consumption appears to “recover” after 48 hours, growth remains impaired. This suggests that, although oxygen demands are being met after exposure, energy resources are still being consumed to regulate the high‐environmental TAmm. As the first study of its kind, this research contributes valuable insight into the effects of elevated ambient ammonia on Nile tilapia under aquaculture conditions. Because Nile tilapia is a relatively new and globally important culture species, there is still much to be learned about optimizing growth of this species under aquaculture conditions. To date, Nile tilapia research has been largely limited to endpoint studies, those which are measured in terms of mortality resulting from temperature and toxicants. Because Nile tilapia is a fish that provides a real solution to the pressing issue of collapsing fisheries stocks, further research into the optimization of growth in this species is clearly warranted The present study has highlighted whole‐body growth effects of ammonia on Nile tilapia. This study has also highlighted the need for a deeper understanding of the physiological mechanisms that become activated when tilapia encounter sub‐lethal conditions of ammonia toxicity, a common obstacle in the culturing of dense populations of fish. This is particularly important in light of previous studies that have demonstrated benefits of improved fish growth condition and productivity in response 53 to elevated ammonia. Previous studies have highlighted the detrimental effects of ammonia on gill tissue. The next step in investigating the effects of TAmm on Nile tilapia growth should be to examine the time course of the impact of elevated ammonia concentrations on gill histology. Specifically, if ammonia is damaging the gill respiratory surfaces how is this disruption being compensated for so that negative effects on the growth potential are not observed at the concentrations examined in this study. It will also be important to determine the relative contributions of each component of total water ammonia (NH3/NH4+) on the growth physiology of tilapia. Furthermore, it is imperative to investigate the specific metabolic profile of tilapia exposed to ammonia concentrations encountered under different growth conditions in order to understand how tilapia regulate this “waste” product. These investigations should include an examination of internal lactate levels in tissues to determine the role of anaerobic metabolism in the response to ammonia exposure in tilapia. In order to optimize Nile tilapia growth in aquaculture, further research is required into internal distribution of ammonia, and its effects on specific tissues. 54 REFERENCES Arillo, A. Margiocco, C. Melodia F. and P. Mesi. 1981. Effects of ammonia on liver
lysosomal functionality in Salmo gairdneri Rich. Journal of Experimental
Zoology 218: 321–326.
Balarin, J.D. and R. D. Haller. 1982. The intensive culture of tilapia in tanks, raceways and cages. In: Recent Advances in Aquaculture Edited by: J.F. Muir and R.J. Roberts, pp. 265–356. Croom Helm Ltd, London. Beamish, F.W.H. and A. Tandler. 1990. Ambient ammonia, diet and growth in lake trout. Aquatic Toxicology 17: 155–156. Beaumont, M., Butler, P. and E. Taylor. 1995. Plasma ammonia concentration in brown trout in soft acidic water and its relationship to decreased swimming performance. Journal of Experimental Biology 198: 2213‐2220. Beaumont, M., Taylor, E. and P. Butler. 2000. The resting membrane potential of white muscle from brown trout (Salmo trutta) exposed to copper in soft, acidic water. Journal of Experimental Biology 203: 2229‐2236. Benli, A., Köksal, G. and A. Özkul. 2008. Sublethal ammonia exposure of Nile tilapia (Oreochromis niloticus L.): Effects on gill, liver and kidney histology. Chemosphere 72: 1355‐1358. 55 Beouf, G. and P. Payan. 2001. How should salinity influence fish growth? Comparative Biochemistry and Physiology 130: 411‐423. Boutilier, R. G., Heming, T. A. and G. K., Iwama. 1984. Physico‐chemical parameters for use in fish respiratory physiology. In: Fish Physiology, vol. 10A Edited by: W. S. Hoar and D. J. Randall pp. 401‐430. New York: Academic Press. Busacker, G. P., Adelman, I. R. and E. M. Goolish. 1990. Growth. In: Methods for fish biology. Edited by: C. B. Schreck and P. B. Moyle. pp. 363‐382 American Fisheries Society, Bcthesda. Maryland. Cech, J. J. Jr . 1990. Respirometry. In: Methods for Fish Biology Edited by: Schreck, C. B. & Moyle, P. B. pp. 335–362. Bethesda, MD: American Fisheries Society. Cheng, S., Lee, W., Shieh, L. and J. Cehn. 2004. Increased production and excretion of urea in the Kuruma shrimp (Marsupenaeus japonicas) exposed to combined environments of increased ammonia and nitrite. Archives of Environmental Contamination and Toxicology 47: 352‐362. Clark, E., Harman, J. and J. Forster. 1984. Production of metabolic and waste products by intensively farmed rainbow trout, Salmo gairdneri Richardson. Journal of Fish Biology 67: 381‐393. Crab, R., Avnimelech, Y., Defoirdt, T., Bossier, P. and W. Verstraete. 2007. Nitrogen removal in aquaculture for a sustainable production, Aquaculture 270: 1‐14. 56 Eddy, F., Kunzlik, P. and R. Bath. 1983. Uptake and loss of nitrite from the blood of Rainbow trout, Salmo gairdneri Richardson, and Atlantic salmon, Salmo salar L. in fresh water and dilute sea water. Journal of Fish Biology 23: 105‐116. Eddy, F. and E. Williams. 1987. Nitrite and freshwater fish. Chemical Ecology 3: 1‐38. Eddy, F. B. 2005. Ammonia in estuaries and effects on fish. Journal of Fish Biology 67:1495‐1513. Edwards, P., Lin, C. and A. Yakupitiyage. 2000. Semi‐intensive pond aquaculture. In: Beveridge. M.C.M. and McAndrew, B. Editors, 2000. Tilapias: Biology and Exploitation, Kluwer Academic Publishing, Dordrecht, pp. 377‐403. El‐Sayed, A.‐F.M, 1999. Alternative dietary protein sources for farmed tilapia, Oreochromis spp., Aquaculture 179: 149–168. El‐Sayed, A.‐F.M. 2006. Tilapia Culture ‐ CABI Publishing, Wallingford El‐Shafaia, S. A., El‐Goharya, F. A., Nasra, F. A., van der Steenb, N. P. and H. J., Gijzenb. 2004. Chronic ammonia toxicity to duckweed‐fed tilapia (Oreochromis niloticus). Aquaculture 232: 117–127. Eng, C., Paw, J. and F. Guarin. 1989. The environmental impact of aquaculture and the effects of pollution on coastal aquaculture development in Southeast Asia. Marine Pollution Bulletin 20: 335‐344. 57 Evans, D. H., Piermarini, P. M. and Choe, K. P. 2005. The multifunctional fish gill: dominant site of gas exchange, osmoregulation, acid‐base regulation, and excretion of nitrogenous waste. Physiological Reviews 85: 97 ‐177. FAO (Food and Agriculture Organization). 2008. The state of the world fisheries and aquaculture 2008. FAO, Rome, Italy. FAO (Food and Agriculture Organization). 2006. The state of the world fisheries and aquaculture 2006. FAO, Rome, Italy. FAO (Food and Agriculture Organization). 2005. Review of state of world marine fishery resources – FAO publication 457. FAO, Rome, Italy. FAO (Food and Agriculture Organization). 2000. The state of the world fisheries and aquaculture 2000. FAO, Rome, Italy. Felipo, V. and R. Butterworth. 2002. Neurobiology of Ammonia. Progress in Neurobiology 67: 259‐279. Fitzsimmons, K. 2006. Prospect and potential for global production. Edited by: C. Lim and C.D. Webster, In: Tilapia biology, Culture and Nutrition, Food Products Press, New York pp. 51–72. Goldstein, L. and R. Forster. 1961. Source of ammonia excreted by the gills of the marine teleost, Myoxocephalus scorpius. American Journal of Physiology 200: 1116‐
1118. 58 Ip, Y. K., Chew, S. F. and D. J., Randall. 2001. Ammonia toxicity, tolerance, and excretion. In: Fish Physiology, vol. 20 Edited by: Wright, P.A., Anderson, P.M., Academic Press, New York, pp. 109–148. Ivancic, I. and D. Deggobis. 1984. An optimal manual procedure for ammonia analysis in natural waters by the indophenol blue method. Water Research 18: 1143–1147. Jenkins. M. 2003. Prospects for biodiversity. Science 302: 1175–1177. Jensen, F. 1990. Nitrite and red cell function in carp: control factors for nitrite entry, membrane potassium ion permeation, oxygen affinity and methaemoglobin formation. Journal of Experimental Biology 152: 149‐166. Jensen, F. 2003. Nitrite disrupts multiple physiological functions in aquatic animals . Comparative Biochemistry and Physiology – Part A: Molecular and Intergrative Physiology 135: 9‐24. Landesman, L. 1994. Negative impacts of coastal tropical aquaculture. World Aquaculture 25: 12‐17. Lemarié, G., Dosdat, A., Covès, D., Dutto, G., Gasset, E. and J.P. Ruyet. 2004. Effect of chronic ammonia exposure on growth of European seabass (Dicentrarchus labrax) juveniles, Aquaculture 229: 479–491. Liao, P. and R. Mayo. 1974. Intensified fish culture combining water reconditioning and pollutant abatement. Aquaculture 3: 61‐85. Linton, T. K., Reid, S. D. and Wood, C. M. 1997. The metabolic costs and physiological 59 consequences to juvenile rainbow trout of a simulated winter warming scenario in the presence or absence of sublethal ammonia. Transactions of the American Fisheries Society 126: 259–272. Linton, T., Morgan, I., Walsh, P. and C. Wood. 1998. Chronic exposure of rainbow trout (Oncorhynchus mykiss) to simulated climate warming and sublethal ammonia: a year‐long study of their appetite, growth and metabolism. Canadian Journal of Fisheries and Aquatic Sciences 55: 576‐586. Lyssenko, C. and F. Wheaton. 2006. Impact of rapid impulse operating disturbances on ammonia removal by tickling and submerged‐upflow biofilters for intensive recirculating aquaculture. Aquacultural Engineering 35: 38‐50. MacLennan, D.N. 1995. Technology in Capture Fisheries. Paper presented at the Government of Japan/FAO International Conference on Sustainable Contribution of Fisheries to Food Security, Kyoto, Japan 4–9 December 1995; and 1997. Madison, B.N., Dhillon, R.S., Tufts, B.L. and Y.X. Wang. 2009. Moderate ammonia promotes growth in walleye (Stizostedion vitreum). Journal of Fish Biology 74: 872‐890. Margiocco, C., Arillo, P., Mensi, P. and G. Schenone. 1983. Nitrite accumulation in Salmo gairdneri rich. and haematological consequences. Aquatic Toxicology 3: 261‐270. McKenzie, D., Randall, D., Lin, H. and S. Aota. 1993. Effects of changes in plasma pH, CO2 and ammonia on ventilation in trout. Fish Physiology and Biochemistry 10: 507‐
515. 60 Meade, J. 1985. Allowable ammonia for fish culture. Progressive Fish Culturist 47: 135‐
145. Mires, D. 1995. The tilapias. In: Nash, C.E., Novotony, A.J. Edited by: Production of Aquatic Animals, Chap. 7. Elsevier, New York, NY, USA, pp. 133–152. Miron, D.S., Moraes, B., Becker, A.G., Crestani, M., Spanevello, R., Loro, V. L and B. Baldissrotto. 2008. Ammonia and pH effects on some metabolic parameters and gill histology of silver catfish, Rhamdia quelen (Heptapteridae). Aquaculture 277: 192‐196. Mommsen, T. and P. Walsh. 1992. Biochemical and environmental perspectives on nitrogen metabolism in fishes. Experientia 48: 583–593. Monfort, P., Kosenko, E., Erceg, S., Canales, J. J. and V. Felipo. 2002. Molecular mechanism of acute ammonia toxicity: role of NMDA receptors. Neurochemistry International 41: 95–102. Mullon, C., Fréon P. and P. Cury. 2005. The dynamics of collapse in world fisheries. Fish and Fisheries 6: 111‐120. Naylor, R.L., Goldburg, R.J., Primavera, J.H., Kautsky, N., Beveridge, M.C., Clay, J., Folke, C., Lubechenco, J., Mooney, H. and M. Troell. 2000. Effect of aquaculture on world fish supplies. Nature. 405: 1017‐1024. Pauly, D., Christensen, V., Dalsgaard, J., Froese, R. and F. Torres Jr. 1998. Fishing down marine food webs. Science 279: 860–863. 61 Pauly, D. and J. Maclean. 2003. In a Perfect Ocean: the State of Fisheries and Ecosystems in the North Atlantic Ocean, Island Press. Person‐Le Ruyet, J., Galland, R., Le Roux, A. and Chartois, H. 1997. Chronic ammonia toxicity in juvenile turbot (Scophthalmus maximus). Aquaculture 154: 155–171. Pillay, T. V. R. and M. N. Kutty. 2005. Aquaculture: Principles and Practices (2nd edition) Blackwell Publishing, Oxford. Popma, T.J. and L.L. Lovshin. 1996. Worldwide prospects for the commercial production of tilapias. International Center for Aquaculture, Research and Development Series 41.
Randall, D. and P. Wright. 1987. Ammonia distribution and excretion in fish. Fish physiology and biochemistry 3: 107‐120. Randall, D., Wilson, J., Peng, K. Kok, T., Kuah, S., Chew, S., Lam, T. and Y. Ip. 1999. The mudskipper, Periophthalmodon schlosseri, actively transports NH4+ against a concentration gradient. American Journal of Physiology – Regulatory Integrative Comparative Physiology 277: 1562‐1567. Randall, D. J. and T. K. N. Tsui. 2002. Ammonia toxicity in fish. Marine Pollution Bulletin 45: 17‐23. Rasmussen, R. S. and B. Korsgaard. 1996. The effect of external ammonia on growth and food utilization of juvenile turbot (Scophthalmus maximus L.). Journal of Experimental Marine Biology and Ecology 205: 35‐48. 62 Read, P. and F. Fernandes. 2003. Management of environmental impacts of marine aquaculture in Europe. Aquaculture 226: 139–163. Redner, B. D. and R.R. Stickney. 1979. Acclimation to Ammonia by Tilapia aurea. Transactions of the American Fisheries Society 108: 383‐388.
Robinette, H. R..1976. Effects of selected sublethal levels of ammonia on the growth of channel catfish Ictalurus punctatus. Progressive Fish Culturist 38: 26–29. Stickney, R. R. 1986. Tilapia, Edited by: R.R. Stickney. In: Culture of Nonsalmonid Freshwater Fishes (pp. 57‐89). Boca Raton, FL: CRC Press. Smart, G. 1976. The effect of ammonia exposure on gill structure of the rainbow trout (Salmo gairdneri). Journal of Fish Biology 8: 471– 475 Smith, H. and G. Rumsey. 1976. Nutrient utilization by fish, In: First Int. Symp., Feed Composition, Animal Nutrient Requirements, and Conputerization of Diets, Edited by: Fonnesbeck, P., Harris, L. and L. Kearl. Utah Agricultural Experiment Station, Utah State University, Logan, Utah, 320. Tidwell, J.H. and G.L, Allan. 2001. Fish as food: aquaculture’s contribution. Ecological and economic impacts and contributions of fish farming and capture fisheries. European Molecular Biology Organization reports 2: 958‐973. Tomasso, J.R. 1994. Toxicity of nitrogenous wastes to aquaculture animals. Reviews in Fisheries Science 2: 291–314. 63 Tovar, A., Moreno, C., Mánuel‐Vez, M. And M. Garcia‐Vargas. 2000. Environmental impacts of intensive aquaculture in marine waters. Water Research 34: 334‐342. Wik, T., Lindon, B. and P. Wramner. 2008. Integrated dynamic aquaculture and wastewater treatment modelling for recirculating aquaculture systems. Aquaculture (In Review). Wilson, R., Wright, P., Munger, S. and C. Wood. 1994. Ammonia excretion in freshwater rainbow trout (Onchorynchus mykiss) and the importance of gill boundry layer acidification: lack of evidence for Na+/NH4+ exchange. Journal of Experimental Biology 191: 37‐58. Wood, C. M. 1988. Acid‐based and ionic exchanges at gills and kidney after exhaustive exercise in the rainbow trout. Journal of Experimental Biology 136: 461‐481. Wood, C. 1993. Ammonia and urea metabolism and excretion. In The Physiology of Fishes. D. Evans (Ed.), pp.379‐425. Boca Raton, CRC Press. Wood, C.M. 2001. Influence of feeding, exercise, and temperature on nitrogen metabolism and excretion. In: Nitrogen Excretion. Edited by: Wright, E., Wright, P., and P. Anderson. San Diego, CA: Acedemic. Wood, C. M. 2004. Dogmas and controversies in the handling of nitrogenous wastes: Is exogenous ammonia a growth stimulant in fish? Journal of Experimental Biology 207: 2043‐2054. 64 Wright, P. and C. Wood. 1985. An analysis of branchial ammonia excretion in the freshwater rainbow trout: Effects of environmental pH change and sodium uptake blockade. Journal of Experimental Biology 114: 329‐353. Wurts, W.A. 2000. Sustainable aquaculture in the twenty‐first century. Reviews in Fisheries Science 8: 131–150. Zieve, L. 1966. Pathogenesis of hepatic coma. Archives of Internal Medicine 118: 211‐
223. 65 APPENDICES APPENDIX 1: Distribution of Series 1 weight data 66 APPENDIX 2: Statistical results from data represented in the figures of Chapter 3: Results Figure Test Test Value p‐value Post hoc test Figure 1 Two‐way ANOVA F 7,441 = 4.6326 p < 0.05 Tukey‐kramer p < 0.05 Figure 2 Two‐way ANOVA F 7,441 = 4.9607 p < 0.05 Tukey‐kramer p < 0.05 Figure 3 Two‐way ANOVA F 18,8406= 3.1168 p < 0.05 Tukey‐kramer p < 0.05 Figure 4 Two‐way ANOVA F 18,8406= 1.8722 p < 0.05 Tukey‐kramer p < 0.05 Figure 5 Two‐way ANOVA F 18,8406= 2.2943 p < 0.05 Tukey‐kramer p < 0.05 Figure 6 Two‐way ANOVA F 5,275 = 5.6236 p < 0.05 Tukey‐kramer p < 0.05 Figure 7 Two‐way ANOVA F 7,441 = 2.2943 p < 0.05 Tukey‐kramer p < 0.05 Figure 8 Two‐way ANOVA F 5,275 = 26.7131 p < 0.05 Tukey‐kramer p < 0.05 Figure 9 F 7,441 = 2.5462 p < 0.05 Tukey‐kramer p < 0.05 F 11,132.1 = 32.6327 p < 0.05 Tukey‐kramer p < 0.05 F 11,132.1 = 34.7608 p < 0.05 Tukey‐kramer p < 0.05 F 11,132.1 = 5.0903 p < 0.05 Tukey‐kramer p < 0.05 Figure 13 Two‐way ANOVA repeated measures
Two‐way ANOVA repeated measures
Two‐way ANOVA repeated measures
Two‐way ANOVA repeated measures
Two‐way ANOVA F 11,132.1 = 2.9658 p < 0.05 Tukey‐kramer p > 0.05 Figure 14 Two‐way ANOVA F 15,1890 = 4.9929 p < 0.05 Tukey‐kramer p < 0.05 Figure 15 Two‐way ANOVA F 15,1890 = 0.7755 p > 0.05 Tukey‐kramer p > 0.05 Figure 10 Figure 11 Figure 12 p‐value 67 APPENDIX 3: Coughing observations Appendix 3: Mean coughing rates of tilapia upon exposure to ammonia (N = 7). Observations occurred over an 8 hour period (one‐way ANOVA, p<0.0001). Significant differences between groups are represented by (*). 68 APPENDIX 4: Tag surgery for juvenile tilapia < 10g (Protocol RM‐T‐004‐2007) Description: Tilapia Surgery protocol Objective: Juvenile Tilapia Surgical set up Outline: Tricaine Methanesulfonate (MS‐222 ) will be used to induce a state of unconsciousness in juvenile tilapia (~10g) as a precursor for surgery. MS‐222 will be prepared at a concentration of 100mg/L with 200mg/L of Sodium bicarbonate in quantities upwards of 2L. The surgical table will consist of a recovery container (2‐4L of fresh aerated water), an adjustable surgical platform (see images, with fluid drain, porous mesh and intubation connection), excess MS‐222, intubation apparatus (beaker with tube and flow regulation attachment) and a syringe (for priming tubes). Fish ~10g will be immersed in the inducing container at concentration 100mg/L MS‐222 for a period of 5‐8 minutes/when no longer responsive to mechanical stimuli (no response to loss of homeostasis). The fish will then be relocated to the surgical platform, the intubation tube placed in its mouth (already flowing) and flow regulated (after intubation flow should be cut and then readjusted for minimal flow where unconscious respiration reoccurs). Following unconsciousness, an incision of ~1‐2cm will be made on the ventral side of the fish, beginning anterior of the anal pore and directed towards the anterior of the fish. Following implantation of PIT tag 1‐2 sutures will be applied as necessary. Post‐operation fish should be allowed to recover for a period of approximately 2 minutes in the recovery chamber. This should be followed by appropriate post‐op protocol. 69 Appendix 4: Image of surgical set up. 70