Global climate change is impacting and will continue to impact on marine and estuarine fish and fisheries. Data trends show climate change effects ranging from fish growth, digestion physiology and performance in marine and freshwater ecosystems. The present study was designed to develop a concept for a cause and effect understanding with respect to climate-induced temperature and salinity changes and to explain ecological findings based on physiological processes. The concept is based on a wide comparison of fish species. The preliminary conclusion can be drawn that warming will cause a shift of distribution limits for fish species with a change in growth performance, gastric evacuation performance and physiology, or even extinction of the species in the world. In association with the elevated seawater temperature growth performance will also be changed with water quality parameters, for example, salinity. Our interpretations of evidence include many uncertainties about the future of affected fish species. Therefore, it is essential to conduct research on the physiology and ecology of marine, estuarine and freshwater fishes, particularly in the tropics where comparatively little research has been conducted and where temperature fluctuation is comparatively lower. As a broader and deeper information base accumulates, researchers will be able to make more accurate predictions and forge relevant solutions.

INTRODUCTION

Global warming may be the most important environmental problem the world faces (Weart 2003; Houghton 2005) and possible impact of generally elevated and extreme temperatures on freshwater and seawater aquatic life are important scientific questions under consideration (Reid et al. 1997). During the last 10–15 years, seawater temperatures throughout much of the globe have changed at unprecedented rates (IPCC 2007; Solomon et al. 2007; Hoegh-Guldberg & Bruno 2010). All organisms have lethal limits to their temperature range (Hokanson 1977) and yet within this range they also have optimal temperatures for development of structure and function (Rombough 1997). Within an ectotherm's tolerance limits, variation in temperature will influence metabolism (Rombough 1997) and therefore related physiological processes, affecting growth (Nicieza & Metcalfe 1997), development (Koumoundouros et al. 2001), and performance encompassing physiological and behavioural capabilities (Fuiman & Higgs 1997). Growth is the most commonly measured response in ectothermic animals and is often measured in isolation as indicative of response to temperature (McMullen & Middaugh 1985; Zhang & Runham 1992). The relative importance of temperature change in the tropics has been alluded to (Rombough 1997; Hunt von Herbing 2002), but rarely tested. Temperature variation of only a few degrees represents a proportionally large change for ectothermic organisms especially fish physiological systems (Feder 1978; Relyea 2002).

It is important to recognize that the observed effects of global warming on fishes at the various levels of biological organization (organismal, population and community ecosystem) result from physiological changes at molecular, cellular and whole organism levels and that the ultimate effects of global warming at the ecosystem level will build on species-specific responses (Portner 2001, 2002). Continuous exposure to elevated temperature can make fish sterile and/or sexually incompetent (Majhi & Das 2013). Species that suffer from stress in nature because of the steady increase in temperature, may be one reason for the lower natural reserves of these species in nature (Dalvi et al. 2009). Such a cause and effect understanding is needed to reliably project the effects of global warming on commercially important fish species and to disentangle these effects from the synergistic effect of fishing pressure on such populations.

This review paper focuses on how global changes (particularly temperature related) impact marine and estuarine fish and fisheries. The amazing aspect of global climate change is the magnitude of the impact of a relatively small temperature change. An increase of a few degrees in atmospheric temperature will not only raise the temperature of the oceans, but also cause major hydrologic changes affecting the physical and chemical properties of water. These will lead to fish, invertebrate and plant species changes in marine and estuarine communities (McGinn 2002). Fishes have evolved physiologically to live within a specific range of environmental variation, and existence outside of that range can be stressful or fatal (Barton et al. 2002). These ranges can coincide for fishes that evolved in similar habitats (Attrill 2002).

Oxygen consumption is often used as an index of metabolism of fishes and is strongly dependent on acclimation temperatures (Majhi & Das 2013). Fish differ in their tolerance to extremes in temperature depending on the species involved, stage of development, environmental temperature dissolved oxygen (DO), pollution, season and extent to which the environment is heated, and that temperature fluctuation affects feeding rate, spawning, DO uptake, pH level and other water quality parameters which would then affect the well-being of the fish.

Temperature of the aquatic environment is important for ensuring survival, distribution and normal metabolism of fish. Failure to adapt to temperature fluctuations is generally ascribed to the inability of fish to respond physiologically with resultant mortality, which is related to changes in the metabolic pathways. Moreover, the temperature changes are complex mixtures of adjustment in hormones, metabolic pathways, catalytic properties of enzymes and behaviour which occur at all functional levels from the molecular and cellular to the whole organism and population (Adeyemo et al. 2003).

This review paper focuses attention on the need for a cause and effect understanding of climate interactions with marine fishes and the ecosystems in which they live. A review is given of broad-scale patterns observed in: (1) temperature tolerance and its effects on fish growth; (2) salinity effects on fish growth; (3) the effects of temperature on fish physiological status; and (4) effects of temperature on gastric evacuation rate of fish.

TEMPERATURE TOLERANCE AND ITS EFFECTS ON FISH GROWTH

Fish generally show temperature optima for growth and survival (Brett 1979; Gadomski & Caddell 1991). These may change with age and size, as juveniles of many species prefer warmer temperatures than adults do (McCauley & Huggins 1979; Pedersen & Jobling 1989). Early life stages may also have different optimal temperatures, which may reflect temporal and spatial field distributions (Gadomski & Caddell 1991; Imsland et al. 1996). Further, the combined effects of size and temperature on growth have been described for several fish species (Brett 1979; Imsland et al. 1996, 2006). The literature is replete with studies that have measured lethal limits, and thus, tolerance to extreme temperatures by fish species (Table 1). These differences appear not only between species but also within species, for example, related to differences in thermal windows due to acclimation or permanent population differences. Measurements of the upper and lower lethal temperatures or critical thermal maxima or minima (Becker & Genoway 1979) of fish species (TCmax and TCmin or TLmax and TLmin) can be used to illustrate the degree of specialization of species and populations on specific thermal environments. Naturally, these thermal endpoints are not static but depend to some degree on acclimation temperature. When a fish species can be acclimated to vastly different temperatures, such as a range of 20 °C, large differences in the upper and lower limits can be observed (Portner & Peck 2010). Within a species, preferred temperatures are often closely related to TCmax (Tsuchida 1995) and often correspond to optimal growth temperatures (Jobling 1981).

Table 1

Temperature window (below or above) for freshwater (F), marine (M), estuarine (E) and anadromous (A) fishes at their different developmental stages (larvae: L, juvenile: J and adult: A) of life

Temperature (°C)
CriticalLethal
SpeciesLife stage (L, J, A)TypeLatitude of field collection (°N or °S)Acclim.Min.Max.Min.Max.Preferred meanReferences
Apogon pacifici 26.5 10.8 35.2    Graham (1971); Mora & Ospina (2001, 2002)  
Plagiotremus azaleus 26.5 13.4 38.2    Mora & Ospina (2001, 2002)  
Cirrhitichthys oxycephalus 26.5 11.4 35.4    Mora & Ospina (2001, 2002)  
Bathygobious ramosus 26.5 12 35.8    Mora & Ospina (2001, 2002)  
Coryphopterus urospilus 26.5 12.3 36    Mora & Ospina (2001, 2002)  
Haemulon steindachneri 26.5 13.2 38.1    Mora & Ospina (2001, 2002)  
Halichoeres dispilus 26.5 15.6 39.7    Mora & Ospina (2001, 2002)  
Thalassoma lucasanum 26.5 16.2 40.1    Mora & Ospina (2001, 2002)  
Malacoctenus zonifer 26.5 12.5 37.3    Mora & Ospina (2001, 2002)  
Lutjanus guttatus 26.5 12 35.9    Mora & Ospina (2001, 2002)  
Eucinostomus gracilis 26.5 12.5 36    Mora & Ospina (2001, 2002)  
Mugil curema 26.5 10.7 34.7    Mora & Ospina (2001, 2002)  
Chromis atrilobata 26.5 11.2 35.3    Mora & Ospina (2001, 2002)  
Stegastes acapulcoensis 26.5 12.6 37.5    Mora & Ospina (2001, 2002)  
Apogon dovii 3 and 9 26.5 13.1 37.8    Mora & Ospina (2001, 2002)  
Apogon novemfasciatus Np 5.5 17–32 15.4 38    Eme & Bennett (2009)  
Bathygobius fuscus Np 5.5 20–35 11.1 41.2    Eme & Bennett (2009)  
Bathygobius sp. Np 5.5 17–34 12.4 41.4    Eme & Bennett (2009)  
Liza vaigiensis Np 5.5 20–35 13 42.2    Eme & Bennett (2009)  
Dascyllus aruanus Np 5.5 17–31 14.3 38    Eme & Bennett (2009)  
Horabagrus brachysoma Np 15–36 15.2 40    Dalvi et al. (2009)  
Apogon maculatus 26–28 13 37.7    Graham (1971)  
Bathygobius ramosus 17–28 9.5 39.5    Graham (1971)  
Bathygobius soporator 26–28 10.1 40.9    Graham (1971)  
Abudefduf saxatilis 26–28 11.6 39.3    Graham (1971)  
Abudefduf troschelii 26.5–29 11.9 39.4    Graham (1971)  
Ambasis commersoni 11.5 28  40.5    Rajaguru & Ramachandran (2001)  
Lates calcarifer 11.5 28  44.5    Rajaguru & Ramachandran (2001)  
Liza dussumeri 11.5 28  44.5    Rajaguru & Ramachandran (2001)  
Etroplus suratensis 11.5 20–35  42.3    Rajaguru & Ramachandran (2001)  
Scatophagus argus 11.5 28  42.5    Rajaguru & Ramachandran (2001)  
Siganus javus 11.5 28  39.5    Rajaguru & Ramachandran (2001)  
Therapon jarbua 11.5 20–35  42    Rajaguru & Ramachandran (2001)  
Ambassis kopsii Np 13.3 22.8 and 26   38.6 38.6  Menasveta (1981)  
Apogon auteus Np 13.3 23–27.3    38.4  Menasveta (1981)  
Chaetodon rostratus Np 13.3 23–29   37.8 37.8  Menasveta (1981)  
Lutianus melabaricus Np 13.3 23–28   37 37  Menasveta (1981)  
Lutianus vitta Np 13.3 23–28   37 37  Menasveta (1981)  
Monocanthus chinensis Np 13.3 22.5–28   35.2 35.2  Menasveta (1981)  
Mugil dussumerii Np 13.3 22.5–29.5   38.4 40.2  Menasveta (1981)  
Plotosus anguillaris Np 13.3 25–28   38.2 38.2  Menasveta (1981)  
Halichocres nigreceus Np 13.3 22.7–28   37.6 35  Menasveta (1981)  
Epinephelus tauvina Np 13.3 22.7–28   37.6 37.6  Menasveta (1981)  
Siganus oramin Np 13.3 22.7–25      Menasveta (1981)  
Sillago sihama Np 13.3 22.8 and 29.5      Menasveta (1981)  
Therapon theraps Np 13.3 22.7      Menasveta (1981)  
Cyprinodon dearborni J and A 18       Brett (1970)  
Ocyurus chrysurus 18 20–32     27.4 Wallace (1977)  
Poecilia sphenops J and A 18       Brett (1970)  
Rivulus marmoratus J and A 18       Brett (1970)  
Haemulon flavolineatum Np 18 27      Sylvester (1973)  
Labeo rohita 18.9 25–35 14 41.3    Chatterjee et al. (2004)  
Cyprinus carpio 18.9 25–35 9.1 41    Chatterjee et al. (2004)  
Anabas testudineus Np 18.9 25–35 13.1 41.1    Sarma et al. (2010)  
Canthigaster jactator J/A 19.4 Np     27 Casterlin & Reynolds (1980)  
Kuhlia sandvicensis J and A 20 23   13.8 34.4  Brett (1970)  
Catla catla 21 30–38 15.2 42.4    Das et al. (2004)  
Pagrus major Np 22 9–28   11 27.8  Woo & Fung (1980)  
Pangasius pangasius 22.6 30–38 14.7 43.5    Debnath et al. (2006)  
Tilapia mossambica Np 25 22–36   16 37.6  Allanson & Noble (1964)  
Xiphophorus maculatus J and A 25.4 15–30 12.9 40.4    Prodocimo & Freire (2001)  
Centropomus undecimalis J and A 26 15–25 12.7  10.1   Shafland & Foote (1983); Howells et al. (1990)  
Sardinops sagax 28 and 34 15.5 and 20.5 6.3 31.3   17.9 Martínez-Porchas et al. (2009)  
Heteropneustes fossilis 29 16 and 29   38.6 30.2 Vasal & Sundararaj (1978)  
Fundulus heteroclitus heteroclitus 30.5 2–34 3.2 38.2    Fangue et al. (2006)  
Atherinops affinis L, J and A 33 14.5–25.5   10.6 31.1 25.2 Doudoroff (1945); Ehrlich et al. (1979)  
Clinocottus analis Np 33 11–23  31.3 27.5 18 Graham (1970)  
Fundulus parvipinnis 33 20     Doudoroff (1945)  
Girella nigricans Np 33 12–28   8.6 30.1  Doudoroff (1942)  
Scomber japonicus J and A 33 16.5   29  Schaefer (1986)  
Sebastes dalli Np 33 13 and 16  25.9   13 Shrode et al. (1982)  
Rypticus nigripinnis 33 17–28 8.6 40    Graham (1971, 1972)  
Hypsoblennius gilberti 34 11–23   6.5 28.8 22.2 Graham (1970)  
Engraulis mordax 34 12–24   10 29.2  Brewer (1976)  
Engraulis mordax J and A 34 8–24   12 33.5  Brewer (1976)  
Citharichthys stigmaeus J and A 34 10–19     11.1 Ehrlich et al. (1979)  
Cheilotrema saturnum J and A 34 17     27.6 Ehrlich et al. (1979)  
Ictalurus punctatus 34.5 20–30 6.3 38.5    Currie et al. (1998)  
Micropterus salmoides 34.5 and 45.8 8–32 7.1 36.9    Fields et al. (1987), Currie et al. (1998)  
Fundulus heteroclitus J and A 37 5–36  38.2  32.3 25 Garside & Chin-Yuen-Kee (1972); Garside & Morrison (1977); Bulger & Tremaine (1985)  
Oncorhynchus mykiss 37 10–20 0.7 29    Currie et al. (1998)  
Gillichthys mirabilis J and A 38 13 or 27  38.2   21 de Vlaming (1971)  
Brevoortia tyrannus 38 7–15   3.8   Lewis (1965)  
Menidia menidia 39 7–28  32.2 5.1 27.3  Brett (1970); Hall et al. (1982)  
Micropogonias undulatus L and J 39     Lankford & Targett (2001)  
Sphoeroides maculatus Np 40 10–28   10.3 30  Brett (1970)  
Lucania parva Np 42 35    37.5  Brett (1970)  
Apeltes quadracus Np 42 25–30    31.5  Brett (1970)  
Gobiesox strumosus Np 42 20    31.5  Brett (1970)  
Tautogolabrus adspersus Np 42 1–22   2.8 27.5  Brett (1970)  
Oncorhynchus tshawytscha 42 20 and 24  25.1   13 Brett (1970)  
Pseudopleuronectes americanus Np 42 7–28   2.7 25.6 18.7 Brett (1970)  
Alosa pseudoharengus 42.4 5–25  31  22.8 Otto et al. (1976)  
Clinocottus globiceps Np 43 21–37    26  Brett (1970)  
Leptocottus armatus Np 43 21–37    29.5  Brett (1970)  
Oligocottus maculosus Np 43 21–37    26.5  Brett (1970)  
Fundulus heteroclitus macrolepidotus 44.3 2–34 3.8 36.4    Fangue et al. (2006)  
Coregonis clupeaformis 44.9 5–22.5    24.5  Edsall & Rottiers (1976)  
Anguilla rostrata J and A 45 5.0–20     18.5 Haro (1991)  
Gobius paganellus Np 45 22–25    31.8  Brett (1970)  
Crenilabrus ocellatus Np 45 22–25    32.8  Brett (1970)  
Mullus barbatus Np 45 22–25    32  Brett (1970)  
Mullus surmuletus Np 45 22–25    30.7  Brett (1970)  
Scorpaena porcus Np 45 22–25    32.5  Brett (1970)  
Box salpa Np 45 22–25    32.5  Brett (1970)  
Sargus vulgaris Np 45 22–25    32.8  Brett (1970)  
Morone saxatilis 45.5 5.0–30   6.9 31.6 24.5–36.1 Coultant et al. (1984); Cook et al. (2006)  
Gadus morhua J and A 47.2      5.5 Despatie et al. (2001)  
Lepidopsetta bilineata J and A 48 6.9–13    24.9  Ames et al. (1978)  
Parophrys vetulus J and A 48 6.9–13    26.1  Ames et al. (1978)  
Oncorhynchus kisutch 49.2 5–23 3.3 24.2    Brett (1952); Konecki et al. (1995)  
Oncorhynchus keta 52 5.0–23 3.8 23    Brett (1952)  
Oncorhynchus nerka 52 5–23 3.1 24.2    Brett (1952)  
Zoarces viviparus Np 54 12  28.8    Zakhartsev et al. (2003)  
Pleuronectes flesus Np 55 12–14    29  Brett (1952)  
Pleuronectes platessa Np 55 12–14    26.5  Jobling (1981)  
Clupea harengus L and A 56 7.5–15   −1.25 20.1  Blaxter (1960); Brawn (1960)  
Pomatoschistus minutus 60 5–17  28.8    Hesthagen (1979)  
Salvelinus alpinus Np 63.5–70 0.5–20  22.5 −1   Baroudy & Elliott (1994); Larsson (2005); Mortensen et al. (2007)  
Pagothenia borchgrevinki Np 78 −19   ≤2.5  Somero & DeVries (1967)  
Trematomus bernacchii Np 78 −19   −2.5  Somero & DeVries (1967)  
Trematomus hansoni Np 78 −19   ≤2.5  Somero & DeVries (1967)  
Temperature (°C)
CriticalLethal
SpeciesLife stage (L, J, A)TypeLatitude of field collection (°N or °S)Acclim.Min.Max.Min.Max.Preferred meanReferences
Apogon pacifici 26.5 10.8 35.2    Graham (1971); Mora & Ospina (2001, 2002)  
Plagiotremus azaleus 26.5 13.4 38.2    Mora & Ospina (2001, 2002)  
Cirrhitichthys oxycephalus 26.5 11.4 35.4    Mora & Ospina (2001, 2002)  
Bathygobious ramosus 26.5 12 35.8    Mora & Ospina (2001, 2002)  
Coryphopterus urospilus 26.5 12.3 36    Mora & Ospina (2001, 2002)  
Haemulon steindachneri 26.5 13.2 38.1    Mora & Ospina (2001, 2002)  
Halichoeres dispilus 26.5 15.6 39.7    Mora & Ospina (2001, 2002)  
Thalassoma lucasanum 26.5 16.2 40.1    Mora & Ospina (2001, 2002)  
Malacoctenus zonifer 26.5 12.5 37.3    Mora & Ospina (2001, 2002)  
Lutjanus guttatus 26.5 12 35.9    Mora & Ospina (2001, 2002)  
Eucinostomus gracilis 26.5 12.5 36    Mora & Ospina (2001, 2002)  
Mugil curema 26.5 10.7 34.7    Mora & Ospina (2001, 2002)  
Chromis atrilobata 26.5 11.2 35.3    Mora & Ospina (2001, 2002)  
Stegastes acapulcoensis 26.5 12.6 37.5    Mora & Ospina (2001, 2002)  
Apogon dovii 3 and 9 26.5 13.1 37.8    Mora & Ospina (2001, 2002)  
Apogon novemfasciatus Np 5.5 17–32 15.4 38    Eme & Bennett (2009)  
Bathygobius fuscus Np 5.5 20–35 11.1 41.2    Eme & Bennett (2009)  
Bathygobius sp. Np 5.5 17–34 12.4 41.4    Eme & Bennett (2009)  
Liza vaigiensis Np 5.5 20–35 13 42.2    Eme & Bennett (2009)  
Dascyllus aruanus Np 5.5 17–31 14.3 38    Eme & Bennett (2009)  
Horabagrus brachysoma Np 15–36 15.2 40    Dalvi et al. (2009)  
Apogon maculatus 26–28 13 37.7    Graham (1971)  
Bathygobius ramosus 17–28 9.5 39.5    Graham (1971)  
Bathygobius soporator 26–28 10.1 40.9    Graham (1971)  
Abudefduf saxatilis 26–28 11.6 39.3    Graham (1971)  
Abudefduf troschelii 26.5–29 11.9 39.4    Graham (1971)  
Ambasis commersoni 11.5 28  40.5    Rajaguru & Ramachandran (2001)  
Lates calcarifer 11.5 28  44.5    Rajaguru & Ramachandran (2001)  
Liza dussumeri 11.5 28  44.5    Rajaguru & Ramachandran (2001)  
Etroplus suratensis 11.5 20–35  42.3    Rajaguru & Ramachandran (2001)  
Scatophagus argus 11.5 28  42.5    Rajaguru & Ramachandran (2001)  
Siganus javus 11.5 28  39.5    Rajaguru & Ramachandran (2001)  
Therapon jarbua 11.5 20–35  42    Rajaguru & Ramachandran (2001)  
Ambassis kopsii Np 13.3 22.8 and 26   38.6 38.6  Menasveta (1981)  
Apogon auteus Np 13.3 23–27.3    38.4  Menasveta (1981)  
Chaetodon rostratus Np 13.3 23–29   37.8 37.8  Menasveta (1981)  
Lutianus melabaricus Np 13.3 23–28   37 37  Menasveta (1981)  
Lutianus vitta Np 13.3 23–28   37 37  Menasveta (1981)  
Monocanthus chinensis Np 13.3 22.5–28   35.2 35.2  Menasveta (1981)  
Mugil dussumerii Np 13.3 22.5–29.5   38.4 40.2  Menasveta (1981)  
Plotosus anguillaris Np 13.3 25–28   38.2 38.2  Menasveta (1981)  
Halichocres nigreceus Np 13.3 22.7–28   37.6 35  Menasveta (1981)  
Epinephelus tauvina Np 13.3 22.7–28   37.6 37.6  Menasveta (1981)  
Siganus oramin Np 13.3 22.7–25      Menasveta (1981)  
Sillago sihama Np 13.3 22.8 and 29.5      Menasveta (1981)  
Therapon theraps Np 13.3 22.7      Menasveta (1981)  
Cyprinodon dearborni J and A 18       Brett (1970)  
Ocyurus chrysurus 18 20–32     27.4 Wallace (1977)  
Poecilia sphenops J and A 18       Brett (1970)  
Rivulus marmoratus J and A 18       Brett (1970)  
Haemulon flavolineatum Np 18 27      Sylvester (1973)  
Labeo rohita 18.9 25–35 14 41.3    Chatterjee et al. (2004)  
Cyprinus carpio 18.9 25–35 9.1 41    Chatterjee et al. (2004)  
Anabas testudineus Np 18.9 25–35 13.1 41.1    Sarma et al. (2010)  
Canthigaster jactator J/A 19.4 Np     27 Casterlin & Reynolds (1980)  
Kuhlia sandvicensis J and A 20 23   13.8 34.4  Brett (1970)  
Catla catla 21 30–38 15.2 42.4    Das et al. (2004)  
Pagrus major Np 22 9–28   11 27.8  Woo & Fung (1980)  
Pangasius pangasius 22.6 30–38 14.7 43.5    Debnath et al. (2006)  
Tilapia mossambica Np 25 22–36   16 37.6  Allanson & Noble (1964)  
Xiphophorus maculatus J and A 25.4 15–30 12.9 40.4    Prodocimo & Freire (2001)  
Centropomus undecimalis J and A 26 15–25 12.7  10.1   Shafland & Foote (1983); Howells et al. (1990)  
Sardinops sagax 28 and 34 15.5 and 20.5 6.3 31.3   17.9 Martínez-Porchas et al. (2009)  
Heteropneustes fossilis 29 16 and 29   38.6 30.2 Vasal & Sundararaj (1978)  
Fundulus heteroclitus heteroclitus 30.5 2–34 3.2 38.2    Fangue et al. (2006)  
Atherinops affinis L, J and A 33 14.5–25.5   10.6 31.1 25.2 Doudoroff (1945); Ehrlich et al. (1979)  
Clinocottus analis Np 33 11–23  31.3 27.5 18 Graham (1970)  
Fundulus parvipinnis 33 20     Doudoroff (1945)  
Girella nigricans Np 33 12–28   8.6 30.1  Doudoroff (1942)  
Scomber japonicus J and A 33 16.5   29  Schaefer (1986)  
Sebastes dalli Np 33 13 and 16  25.9   13 Shrode et al. (1982)  
Rypticus nigripinnis 33 17–28 8.6 40    Graham (1971, 1972)  
Hypsoblennius gilberti 34 11–23   6.5 28.8 22.2 Graham (1970)  
Engraulis mordax 34 12–24   10 29.2  Brewer (1976)  
Engraulis mordax J and A 34 8–24   12 33.5  Brewer (1976)  
Citharichthys stigmaeus J and A 34 10–19     11.1 Ehrlich et al. (1979)  
Cheilotrema saturnum J and A 34 17     27.6 Ehrlich et al. (1979)  
Ictalurus punctatus 34.5 20–30 6.3 38.5    Currie et al. (1998)  
Micropterus salmoides 34.5 and 45.8 8–32 7.1 36.9    Fields et al. (1987), Currie et al. (1998)  
Fundulus heteroclitus J and A 37 5–36  38.2  32.3 25 Garside & Chin-Yuen-Kee (1972); Garside & Morrison (1977); Bulger & Tremaine (1985)  
Oncorhynchus mykiss 37 10–20 0.7 29    Currie et al. (1998)  
Gillichthys mirabilis J and A 38 13 or 27  38.2   21 de Vlaming (1971)  
Brevoortia tyrannus 38 7–15   3.8   Lewis (1965)  
Menidia menidia 39 7–28  32.2 5.1 27.3  Brett (1970); Hall et al. (1982)  
Micropogonias undulatus L and J 39     Lankford & Targett (2001)  
Sphoeroides maculatus Np 40 10–28   10.3 30  Brett (1970)  
Lucania parva Np 42 35    37.5  Brett (1970)  
Apeltes quadracus Np 42 25–30    31.5  Brett (1970)  
Gobiesox strumosus Np 42 20    31.5  Brett (1970)  
Tautogolabrus adspersus Np 42 1–22   2.8 27.5  Brett (1970)  
Oncorhynchus tshawytscha 42 20 and 24  25.1   13 Brett (1970)  
Pseudopleuronectes americanus Np 42 7–28   2.7 25.6 18.7 Brett (1970)  
Alosa pseudoharengus 42.4 5–25  31  22.8 Otto et al. (1976)  
Clinocottus globiceps Np 43 21–37    26  Brett (1970)  
Leptocottus armatus Np 43 21–37    29.5  Brett (1970)  
Oligocottus maculosus Np 43 21–37    26.5  Brett (1970)  
Fundulus heteroclitus macrolepidotus 44.3 2–34 3.8 36.4    Fangue et al. (2006)  
Coregonis clupeaformis 44.9 5–22.5    24.5  Edsall & Rottiers (1976)  
Anguilla rostrata J and A 45 5.0–20     18.5 Haro (1991)  
Gobius paganellus Np 45 22–25    31.8  Brett (1970)  
Crenilabrus ocellatus Np 45 22–25    32.8  Brett (1970)  
Mullus barbatus Np 45 22–25    32  Brett (1970)  
Mullus surmuletus Np 45 22–25    30.7  Brett (1970)  
Scorpaena porcus Np 45 22–25    32.5  Brett (1970)  
Box salpa Np 45 22–25    32.5  Brett (1970)  
Sargus vulgaris Np 45 22–25    32.8  Brett (1970)  
Morone saxatilis 45.5 5.0–30   6.9 31.6 24.5–36.1 Coultant et al. (1984); Cook et al. (2006)  
Gadus morhua J and A 47.2      5.5 Despatie et al. (2001)  
Lepidopsetta bilineata J and A 48 6.9–13    24.9  Ames et al. (1978)  
Parophrys vetulus J and A 48 6.9–13    26.1  Ames et al. (1978)  
Oncorhynchus kisutch 49.2 5–23 3.3 24.2    Brett (1952); Konecki et al. (1995)  
Oncorhynchus keta 52 5.0–23 3.8 23    Brett (1952)  
Oncorhynchus nerka 52 5–23 3.1 24.2    Brett (1952)  
Zoarces viviparus Np 54 12  28.8    Zakhartsev et al. (2003)  
Pleuronectes flesus Np 55 12–14    29  Brett (1952)  
Pleuronectes platessa Np 55 12–14    26.5  Jobling (1981)  
Clupea harengus L and A 56 7.5–15   −1.25 20.1  Blaxter (1960); Brawn (1960)  
Pomatoschistus minutus 60 5–17  28.8    Hesthagen (1979)  
Salvelinus alpinus Np 63.5–70 0.5–20  22.5 −1   Baroudy & Elliott (1994); Larsson (2005); Mortensen et al. (2007)  
Pagothenia borchgrevinki Np 78 −19   ≤2.5  Somero & DeVries (1967)  
Trematomus bernacchii Np 78 −19   −2.5  Somero & DeVries (1967)  
Trematomus hansoni Np 78 −19   ≤2.5  Somero & DeVries (1967)  

Available knowledge indicates that thermal windows are narrow in early life stages, due to developmental constraints and insufficient capacity of central organs in the larvae (Portner et al. 2006) and widen in juveniles and young adults in line with rising performance capacity at small body size. Larger individuals then become more thermally sensitive, due to progressively falling oxygen supply capacity in relation to demand (Portner & Farrell 2008). Sensitivity to cold appears as a very important characteristic in shaping community composition. Low temperatures during winter may increase mortality, either because temperatures fall outside the thermal window or because energy reserves become limiting, especially in smaller individuals that have relatively fewer reserves compared to larger conspecifics (Post & Evans 1989; Sogard 1997). Extreme winter events cause reductions in species abundance and ecosystem changes, for example, in the German Wadden Sea (Woodhead 1964). The consequences of different thermal windows for growth and spawning productivity can be found in the oscillations between Engraulis japonicus and Sebastes melanostictus in the Pacific Ocean (Takasuka et al. 2007, 2008).

The Atlantic cod (Gadus morhua) populations between the southern North Sea and the Arctic North Atlantic display different thermal windows of growth (Portner et al. 2001, 2008). Productivity will also be influenced by the effect of temperature on growth rate (Brander 1995; Teal et al. 2008). An increase in juvenile growth as well as an increase in temperature may result in a decrease in the length and age at first maturation, affecting the growth of adults as surplus energy is channelled into reproduction at an earlier age and smaller size (Heino et al. 2002). The data available for E. japonicus and S. melanostictus in the Japan Sea indicate that recruitment processes and growth occur within the same species-specific range of temperatures (Takasuka et al. 2007, 2008).

A downshift in temperature optimum with increasing size, that is, ontogenetic shift, has been demonstrated in Atlantic cod G. morhua L. (Pedersen & Jobling 1989), plaice Pleuronectes platessa L. (Fonds et al. 1992), turbot Scophthalmus maximus L. (Imsland et al. 1996), halibut Hippoglossus hippoglossus L. (Jonassen et al. 1999) and spotted wolffish Anahichas minor (Imsland et al. 2006). In contrast, no shift in temperature optimum with increasing size was found in brown trout Salmo trutta L. (Elliott 1975), nor in sockeye salmon Oncorhynchus nerka (Brett et al. 1969). Recent experiments with Atlantic salmon parr have indicated temperature optima around 18–19 °C for 4–12 g fish (Forseth et al. 2001; Jonsson et al. 2001), whereas Handeland et al. (2003) reported 13 °C as temperature optima for 40–60 g Atlantic salmon smolts. Growth rate, food intake and feed efficiency ratio of juvenile Atlantic salmon smolts were significantly influenced by temperature and fish size. Increased growth at elevated temperatures agrees with previous studies on Atlantic salmon (Solbakken et al. 1994; Handeland et al. 2000, 2003). In brown trout reared in freshwater, a decrease in appetite has been observed when temperature exceeds 18 °C (Brett 1979). Handeland et al. (2000) reported a linear relationship between specific growth rate (SGR) and temperature in Atlantic salmon smolts in seawater between 4.6 and 14.4 °C, while a rapid decrease in growth rate was observed when temperature reached 18.9 °C.

Several authors have observed maximal growth efficiency within a range of optimal water temperatures, with declining efficiency at both lower and higher temperatures (Björnsson & Tryggvadóttir 1996); however, Wurtsbaugh & Davis (1977) found no temperature effect on growth efficiency when fish were being fed high ration levels. But in many species, digestion is associated with selection of a higher body temperature (Peterson et al. 1993; Dorcas et al. 1997) and it is possible that the postprandial thermophilic response enables a more efficient and/or faster digestion. Teleologically, the existence of a postprandial thermophilic response suggests that the optimal temperature for digestion is higher than optimal temperatures for other behaviours and functions (Wang et al. 2003).

In the tropical fish species, a small variation in temperature resulted in a large variation in growth performance. A 3 °C reduction in rearing temperature decreased the growth, developmental rate and swimming speed in Amphiprion melanopus larvae (Green & Fisher 2004). Growth is a function of cellular activities, which are dictated by general physical laws. Therefore, when temperature-induced changes in growth are standardized for the degree of temperature change, similar patterns could occur across thermal ranges. The growth co-efficient (Q10) values of different fishes are shown in Table 2.

Table 2

Q10 values for growth (mm day−1) of fishes reflecting the effect of temperature on the rate of a given process

SpeciesClimateTemperature change (°C)Q10*Reference
Amphiprion melanopus trop 25–28 2.4 Green & Fisher (2004)  
Channa striatus trop/sub-trop 21.7–27 1.91 Qin & Fast (1998)  
Achirus lineatus sub-trop 24–28 1.66 Houde (1974)  
Anchoa mitchilli sub-trop 28–30 0.10 Houde (1974)  
Anchoa mitchilli sub-trop 24–28 1.67 Houde (1974)  
Archosargus rhomboidalis sub-trop 26–30 0.99 Houde (1974)  
Hippoglossus hippoglossus temp 5–8 5.3 Galloway et al. (1998)  
Clupea harengus temp 10–8 0.8 McGurk (1984)  
Clupea harengus temp 8–6 1.25 McGurk (1984)  
Menidia menidia temp 17–28 4.03 Yamahira & Conover (2002)  
Menidia menidia temp 17–28 4.37 Yamahira & Conover (2002)  
Anarhichas minor (eggs only) temp 6–8 2.38 Hansen & Falk-Petersen (2001)  
Anarhichas minor (eggs only) temp 4–6 5.15 Hansen & Falk-Petersen (2001)  
Morone Americana temp fw 17–21 2.68 Margulies (1989)  
Morone Americana temp fw 13–17 6.42 Margulies (1989)  
Perca fluviatilis temp fw 15–20 4.79 Wang & Eckmann (1994)  
Hybrid Lepomis cyanellus × L. macrochirus temp fw 21–24 2.17 Mischke et al. (2001)  
Hybrid Lepomis cyanellus × L. macrochirus temp fw 19–21 0.95 Mischke et al. (2001)  
Colossoma macropomum trop 20–30 2.97 Saint-Paul (1983)  
Oreochromis mossambicas trop 19–28 2.74 Caulton (1978)  
Protopterus aethiopicus trop 20–30 3.30 This study 
Scophthalmus maximus temp 6–22 3.06 Mallekh & Lagardere (2002)  
Gambusia affinis temp 10–30 2.24 Cech et al. (1985)  
Anguilla anguilla temp 20–27 2.48 Degani et al. (1989)  
Anguilla rostrata temp 15–25 3.67 Degani & Gallagher (1985)  
Cyprinus carpio temp 10–30 2.60 Beamish (1964)  
Carassius auratus temp 10–30 2.29 Beamish & Mookherjii (1964)  
Catostomus commersonii temp 10–30 2.44 Beamish (1964)  
Oncorhynchus nerka temp 5–20 2.02 Brett & Glass (1973)  
Salvelinus fontinalis temp 10–20 2.84 Beamish (1964)  
Pleuronectes platessa temp 2–22 2.06 Fonds et al. (1992)  
Platichthys flesus temp 2–22 2.24 Fonds et al. (1992)  
SpeciesClimateTemperature change (°C)Q10*Reference
Amphiprion melanopus trop 25–28 2.4 Green & Fisher (2004)  
Channa striatus trop/sub-trop 21.7–27 1.91 Qin & Fast (1998)  
Achirus lineatus sub-trop 24–28 1.66 Houde (1974)  
Anchoa mitchilli sub-trop 28–30 0.10 Houde (1974)  
Anchoa mitchilli sub-trop 24–28 1.67 Houde (1974)  
Archosargus rhomboidalis sub-trop 26–30 0.99 Houde (1974)  
Hippoglossus hippoglossus temp 5–8 5.3 Galloway et al. (1998)  
Clupea harengus temp 10–8 0.8 McGurk (1984)  
Clupea harengus temp 8–6 1.25 McGurk (1984)  
Menidia menidia temp 17–28 4.03 Yamahira & Conover (2002)  
Menidia menidia temp 17–28 4.37 Yamahira & Conover (2002)  
Anarhichas minor (eggs only) temp 6–8 2.38 Hansen & Falk-Petersen (2001)  
Anarhichas minor (eggs only) temp 4–6 5.15 Hansen & Falk-Petersen (2001)  
Morone Americana temp fw 17–21 2.68 Margulies (1989)  
Morone Americana temp fw 13–17 6.42 Margulies (1989)  
Perca fluviatilis temp fw 15–20 4.79 Wang & Eckmann (1994)  
Hybrid Lepomis cyanellus × L. macrochirus temp fw 21–24 2.17 Mischke et al. (2001)  
Hybrid Lepomis cyanellus × L. macrochirus temp fw 19–21 0.95 Mischke et al. (2001)  
Colossoma macropomum trop 20–30 2.97 Saint-Paul (1983)  
Oreochromis mossambicas trop 19–28 2.74 Caulton (1978)  
Protopterus aethiopicus trop 20–30 3.30 This study 
Scophthalmus maximus temp 6–22 3.06 Mallekh & Lagardere (2002)  
Gambusia affinis temp 10–30 2.24 Cech et al. (1985)  
Anguilla anguilla temp 20–27 2.48 Degani et al. (1989)  
Anguilla rostrata temp 15–25 3.67 Degani & Gallagher (1985)  
Cyprinus carpio temp 10–30 2.60 Beamish (1964)  
Carassius auratus temp 10–30 2.29 Beamish & Mookherjii (1964)  
Catostomus commersonii temp 10–30 2.44 Beamish (1964)  
Oncorhynchus nerka temp 5–20 2.02 Brett & Glass (1973)  
Salvelinus fontinalis temp 10–20 2.84 Beamish (1964)  
Pleuronectes platessa temp 2–22 2.06 Fonds et al. (1992)  
Platichthys flesus temp 2–22 2.24 Fonds et al. (1992)  

All other values were calculated following Q10 = [R2/R1]10/(T2T1), where T1 and T2 are the temperatures over which the change was recorded, R1 is the rate of a process at T1 and R2 is the rate of the process at T2.

*Denote Q10 values that were published in the cited reference.

The growth of salmonids tends to be optimal at 12–17 °C (Brett 1971; Kosekla et al. 1997a), however they maintain feeding and some growth at temperatures approaching 0°C (Kosekla et al. 1997b). Salmonid digestive processes are influenced by temperature (Bendiksen et al. 2003). Salmonids tend to also show increased macronutrient digestibility with increases in temperature (Atherton & Aitken 1970; Bendiksen et al. 2003). However, sablefish reside in cooler waters and may respond better to cooler temperatures compared to salmonids (Pace 2013).

Some authors have reported that gross conversion efficiency (GCE) increases with decreasing temperature and have suggested that this is an adaptive physiological characteristic of these predominantly cold-water species (Peck et al. 2003). However, other authors have reported that GCE is higher at elevated temperatures (e.g., 13 vs. 2 °C) (Purchase & Brown 2001). The reason why GCE was insensitive (Atlantic cod) or fell (haddock) substantially when temperature was lowered from 11 to 2 °C was unknown (Perez-Casanova et al. 2009), but it is unlikely to be related to food consumption (FC) since Peck et al. (2003) showed that GCE (calculated as GCE = SGR/FC) is higher at lower temperatures, irrespective of FC.

Evidence that warmer temperatures may be beneficial for gut function is also found in studies on warm bodied fishes. Tuna and the lamnid sharks have evolved counter current heat exchange mechanisms for conserving metabolic heat and raising their body temperatures above the ambient temperature (Carey et al. 1971). In both the bluefin tuna (Thunnus thynnus) and the white shark (Carcharodon carcharias), the gut is thermally isolated by a circulatory heat exchange system, allowing it to maintain gut temperatures as much as 10–15 °C above ambient temperature (Stevens & McLeese 1984; Goldman 1997). In the bluefin tuna, the elevated gut temperatures allow the fish to digest a meal up to three times quicker than normal, enabling them to consume and process about three times as much food per day (Stevens & McLeese 1984). Bluefin tuna are also able to slowly increase the temperature of the stomach after a meal, and slowly decrease it when unfed (Stevens & McLeese 1984).

SALINITY EFFECTS ON FISH GROWTH

Development and growth (continuous in fish) are controlled by ‘internal factors’ including endocrinological and neuroendocrinological systems. Among vertebrates, they also are highly dependent on environmental conditions. Beside temperature, salinity is another important and relevant variable in the study of physiology. The combined effect of temperature and salinity somewhat determine the metabolism rate of the organisms and consequently, the extent of distribution of the species (Vernberg & Vernberg 1972). A knowledge of the influence of temperature and salinity on the physiological rates of fish is essential for the interpretation of studies on production in natural environments.

Among other factors, many studies have reported an influence of water salinity together with temperature on fish development and growth. In most species, egg fertilization and incubation, yolk sac resorption, early embryogenesis, swim bladder inflation, larval growths are dependent on salinity. In larger fish, salinity is also a key factor in controlling growth. Temperature and salinity have complex interactions (Gilles & Patrick 2001). All these facts must be considered when developing fish culture (Boeuf & Payan 2001), which essentially aims to produce fish of the best quality at the most economical cost.

The responses of teleost fishes to different rearing salinities affect growth and, possibly, survival (Johnson & Katavic 1986; Lee & Menu 1986; Banks et al. 1991; Murashige et al. 1991; Bone et al. 1995). Many authors have demonstrated the influence of external salinity on growth capacities in fish (see Table 3). Fish adaptive capacity to different salinities depends on the integrated osmoregulatory function of numerous organs, mainly the gills, digestive tract and kidney. Fish gill undergoes marked morphological changes particularly in mitochondria-rich cells (chloride cells) related to its roles in active ion transport (Perry 1997; Evans et al. 1999). Fish have prolactin (PRL) cells which are directly osmosensitive (Grau et al. 1994), and the development and activity of PRL cells are influenced by changes in environmental osmolarity in the cyprinodont fish, Cynolebias whitei (Ruijter et al. 1984).

Table 3

Acclimation to salinity and its influence on growth

SpeciesDrinking rate (ml kg−1 h−1)Reference
Salmo salar (Atlantic salmon) FW 0.1 Fuentes & Eddy (1997)  
SW 3.88 
Salmo salar 6 h SW 2.4 Usher et al. (1988)  
10d SW 7.94 
Oncorhynchus mykiss (rainbow trout)  5.38 Shehadeh & Gordon (1969)  
Scophthalmus maximus (turbot)  1.4 Carroll et al. (1994)  
Limanda limanda (dab)  2.9 Carroll et al. (1994)  
Pleuronectes platessa (plaice)  2.48 
Anguilla anguilla (eel)  1.0 Perrott et al. (1992)  
Merlangius merlangus (whiting)  1.8 Perrott et al. (1992)  
Myxocephalus scorpius (sea scorpion)  7.8 Perrott et al. (1992)  
Agonus cataphractus (pogge)  2.21 Perrott et al. (1992)  
Anarhichas lupus (wolffish)  2.24 Perrott et al. (1992)  
Ammodytes lanceolatus (sand eel)  2.96 Perrott et al. (1992)  
SpeciesDrinking rate (ml kg−1 h−1)Reference
Salmo salar (Atlantic salmon) FW 0.1 Fuentes & Eddy (1997)  
SW 3.88 
Salmo salar 6 h SW 2.4 Usher et al. (1988)  
10d SW 7.94 
Oncorhynchus mykiss (rainbow trout)  5.38 Shehadeh & Gordon (1969)  
Scophthalmus maximus (turbot)  1.4 Carroll et al. (1994)  
Limanda limanda (dab)  2.9 Carroll et al. (1994)  
Pleuronectes platessa (plaice)  2.48 
Anguilla anguilla (eel)  1.0 Perrott et al. (1992)  
Merlangius merlangus (whiting)  1.8 Perrott et al. (1992)  
Myxocephalus scorpius (sea scorpion)  7.8 Perrott et al. (1992)  
Agonus cataphractus (pogge)  2.21 Perrott et al. (1992)  
Anarhichas lupus (wolffish)  2.24 Perrott et al. (1992)  
Ammodytes lanceolatus (sand eel)  2.96 Perrott et al. (1992)  

Data are indicated for different fish species in ml kg−1 h−1 in FW and SW.

They also possess chemoreceptors, situated in the pseudobranch (Rahim et al. 2014), providing information on water salinity. These are connected to the central nervous system and participate in triggering water drinking in seawater. Several hormones are known to be involved in such phenomena, among them the renin angiotensin system (Perrott et al. 1992), and growth hormones (Fuentes & Eddy 1997) are important.

The extra costs for ionic regulation may reduce energy available for growth unless the fish can compensate by increasing its feeding rate (Wootton 1995). Larvae of grey mullet (Mugil cephalus) obtained maximum growth at 22–23 psu (Lee & Menu 1986) and at 16 psu (Murashige et al. 1991) after 15 days of rearing. For seabass (Dicentrarchus labrax), Johnson & Katavic (1986) reported maximum growth at 10–20 psu after 18 days of rearing.

In terms of the influence of salinity on the growing capacities in larger fish, juveniles or adults, data from numerous studies are available and these results are summarized in Table 4. This has been demonstrated for both ‘true’ marine species (i.e., cod or freshwater (FW) species, i.e., carp).

Table 4

Salinity and growth

SpeciesToleranceBest growthReference
Pomatomus saltatrix (Bluefish) =5 and 25 psu  Buckel et al. (1995)  
Micropogonias furnieri (Croaker) +10–30 17–19 Aristizabal Abud (1992)  
Sparus sarba (Sea bream) +0–35 15 Woo & Kelly (1995)  
Oreochromis niloticus (Tilapia) +0–16 Likongwe et al. (1996)  
Oreochromis spilurus (Tilapia) =0–37  Jonassen et al. (1997)  
Oreochromis aureus (Tilapia) =0–27  Chervinsky & Yashouv (1971)  
O. mossambicus (Tilapia) 120 psu 17.5 Suresh & Lin (1992)  
O. niloticus (Tilapia) 36 5–10 Suresh & Lin (1992)  
O. aureus (Tilapia) 40 10–15 Suresh & Lin (1992)  
O. spilurus (Tilapia) 36 Suresh & Lin (1992)  
Hybrid Red (Tilapia) 35 30–35 Suresh & Lin (1992)  
Morone chrysops (White bass) +0 20 psu 0–12 Heyward et al. (1995)  
Chanos chanos (Milkfish) +0 55 55 Swanson (1998)  
Chanos chanos (Milkfish) +0 35 Alava (1998)  
Trinectes maculatus (Hogchoker) +0 30 30 Peters & Boyd (1972)  
Trinectes maculatus (Hogchoker) +0 15  Peterson-Curtis (1997)  
Gadus morhua (Cod) 14 and 28 psu 14 Dutil et al. (1997)  
Mugil sp. (Mullet) +3–24 17 Peterson et al. (1999b)  
Chelon labrosus (Grey mullet) =(5–25)  Cardona & Castello-Orvay (1997)  
Salaria fluiatilis (Blenny) +0–35 12 Plaut (1999)  
Micropogonias undulatus (Atlantic croaker) +0–20 Peterson et al. (1999a)  
Sparus aurata (Gilthead seabream) +8–38 28 Conides et al. (1997)  
Lutjanus argentiventris (Amarillo snapper) +3–24 psu 30 Serrano-Pinto & Caravea-Patino (1999)  
Oncorhynchus mykiss (Rainbow trout) +0–16 Morgan & Iwama (1991)  
Salvelinus alpinus (Arctic charr) +0–35 0–20 Arnesen et al. (1993)  
Scophthalmus maximus (Turbot) +0–35 10–19 Gaumet et al. (1995)  
SpeciesToleranceBest growthReference
Pomatomus saltatrix (Bluefish) =5 and 25 psu  Buckel et al. (1995)  
Micropogonias furnieri (Croaker) +10–30 17–19 Aristizabal Abud (1992)  
Sparus sarba (Sea bream) +0–35 15 Woo & Kelly (1995)  
Oreochromis niloticus (Tilapia) +0–16 Likongwe et al. (1996)  
Oreochromis spilurus (Tilapia) =0–37  Jonassen et al. (1997)  
Oreochromis aureus (Tilapia) =0–27  Chervinsky & Yashouv (1971)  
O. mossambicus (Tilapia) 120 psu 17.5 Suresh & Lin (1992)  
O. niloticus (Tilapia) 36 5–10 Suresh & Lin (1992)  
O. aureus (Tilapia) 40 10–15 Suresh & Lin (1992)  
O. spilurus (Tilapia) 36 Suresh & Lin (1992)  
Hybrid Red (Tilapia) 35 30–35 Suresh & Lin (1992)  
Morone chrysops (White bass) +0 20 psu 0–12 Heyward et al. (1995)  
Chanos chanos (Milkfish) +0 55 55 Swanson (1998)  
Chanos chanos (Milkfish) +0 35 Alava (1998)  
Trinectes maculatus (Hogchoker) +0 30 30 Peters & Boyd (1972)  
Trinectes maculatus (Hogchoker) +0 15  Peterson-Curtis (1997)  
Gadus morhua (Cod) 14 and 28 psu 14 Dutil et al. (1997)  
Mugil sp. (Mullet) +3–24 17 Peterson et al. (1999b)  
Chelon labrosus (Grey mullet) =(5–25)  Cardona & Castello-Orvay (1997)  
Salaria fluiatilis (Blenny) +0–35 12 Plaut (1999)  
Micropogonias undulatus (Atlantic croaker) +0–20 Peterson et al. (1999a)  
Sparus aurata (Gilthead seabream) +8–38 28 Conides et al. (1997)  
Lutjanus argentiventris (Amarillo snapper) +3–24 psu 30 Serrano-Pinto & Caravea-Patino (1999)  
Oncorhynchus mykiss (Rainbow trout) +0–16 Morgan & Iwama (1991)  
Salvelinus alpinus (Arctic charr) +0–35 0–20 Arnesen et al. (1993)  
Scophthalmus maximus (Turbot) +0–35 10–19 Gaumet et al. (1995)  

The influence of salinity on growth of different species of fish are indicated as follows: (positive + or neutral=), tolerance (maximum salinity or range tested), the best salinity conditions for growth and the reference.

In sea bream (Sparus aurata) larvae, growth was estimated at 15–40 psu and the best results, in terms of weight increase and swimbladder inflation, were recorded at 25 psu (Tandler et al. 1995). In greenback flounder (Rhombosolea tapirina), egg fertilization, incubation and yolk sac resorption are dependent on salinity, with 28 psu resulting in the best overall performance (Hart & Purser 1995). In other flatfish, such as the summer flounder (Paralichthys dentatus) or southern flounder (P. lethostigma), early development and larval growth were also affected by salinity, optimal conditions being 8–14 and 5–30 psu, respectively (Smith et al. 1999). For mulloway (Argyrosomus japonicas) and milkfish (C. chanos), yolk resorption, early embryogenesis and larval growth were optimal at salinities of 5–12.5 and 20–35 psu (Fielder & Bardsley 1999). In striped bass (Morone saxatilis), growth and yolk sac utilization were optimal at 5 psu, compared with 0 and 10 psu (Peterson et al. 1996). In the red tilapia (hybrid Oreochromis mossambicus × O. urolepis), early development and growth were optimal at 18 psu (tested range, 4–36 psu) (Watanabe et al. 1989). A change from 0 to 36.6 psu saltwater (SW) did not affect growth in the tilapia O. spilurus when the salinity was increased progressively over 120 h (Jonassen et al. 1997). In the salmonid Oncorhynchus keta (chum salmon), increasing the salinity to 33.5 psu during rearing, following 7 weeks in FW (7–10 g fish), resulted in increased growth (Kojima et al. 1993). Salmonids can grow better in SW during the winter in net-pen culture, often independently of the higher salinity, since temperature is higher than in rivers during this season. Also depending on the duration of the experiment, or on the developmental stage of the animal, results may differ between authors. This was demonstrated for smoltifying salmonids, where the smolt status profoundly influenced salinity ‘receptivity’, osmotic capacities and growth (Boeuf 1993). The metabolic rate of newly hatched steelhead trout (O. mykiss) alevins was significantly lower in 8 psu water, and higher at 12, in comparison to either 0 or 4 psu (Morgan et al. 1992).

In experimenting with FW species, carp Cyprinus carpio, white Amur Ctenopharyngodon idella and juvenile Russian sturgeon Acipenser guldenstaedti, it was shown that a salinity of 2 psu considerably increased growth rate and food efficiency, by improving the food conversion rate (Konstantinov & Martynova 1993). Moreover, respiration rate and MO2 decrease, and fluctuations between 0 and 2 psu are even more efficient. In contrast to this, in ‘true’ SW species, such as cod G. morhua or turbot Scophthalmus maximus, the growth rate is significantly increased at intermediate salinity conditions of 19 psu (Lambert et al. 1994; Gaumet et al. 1995; Dutil et al. 1997; Imsland et al. 2001). In cod, an increase in food conversion efficiency results in a higher growth rate at lower salinity, whereas turbot increase their food intake (Gaumet et al. 1995). Imsland et al. (2001) concludes there is better growth and better food efficiency if turbot is reared at intermediate salinities (15–19) psu. In conclusion, we can say that if fish growth is obviously influenced by the water temperature, salinity interactions also occur.

EFFECTS OF TEMPERATURE ON FISH PHYSIOLOGICAL STATUS

Studies of blood parameters have proven to be a valuable approach for analysing the health status of fish and help in understanding the relationship of blood characteristics to the habitat and adaptability of species to the environment (Bahmami et al. 2001; Fazio et al. 2012). The changes in levels of fish blood parameters, such as red blood cell (RBC), white blood cell (WBC), haemoglobin (Hb) and haematocrit (Hct), will give an insight into a fish's health status (Harikrishnan et al. 2011). In the aquatic habitat, the fish homeostatic system is continuously affected by the changes of the level of temperature concentration (Imsland et al. 2008). It is important for cells to maintain homeostasis for the organism to remain healthy and fish could mediate physiological efficiency to compensate change induced by temperature (Prophete et al. 2006). Optimum temperature has been reported to enhance immune responses in fish whereas lower temperatures adversely affected immune function (Bly & Clem 1992). Temperature is known to affect the enzyme reaction, immune response, haematological parameters and plasma electrolytes (Tanck et al. 2000). In recent years haematological indices have been used to determine the effect of stress (Wedemeyer & Yasutake 1977). In fish, cold temperatures increase the oxygen requirement, cardiac output and blood flow (Julian et al. 1989) and in common carp (Cyprinus carpio) haemolysis has been reported during acute water temperature changes from 8 to 4 °C (Chen et al. 1995).

Haemoglobin is a conjugated protein responsible for carrying oxygen in fish body, and its concentration is closely related to RBC counts (Clark et al. 1979). Haemoglobin concentration and RBC count of sheatfish (Silurus soldatovi) increased at first and later decreased when temperature ranged between 4 and 30 °C (Zhi et al. 2008). An increase in the WBC count was found in rainbow trout (Oncorhynchus mykiss) (Houston et al. 1996) and carp (Engelsma et al. 2003) at high temperature and after cold shock, respectively. Ndong et al. (2007) found that WBC counts decreased significantly when Mozambique tilapias (O. mossambicus) were transferred from 27 to 19 or 35 °C. Tilapias stop feeding when temperature goes below 15 °C, and are unable to reproduce below 20 °C (Sun et al. 1995). High water temperature may pose physiological stress in tilapia, thereby causing cellular injury in kidney and liver as well as reducing the level of erythropoietin, thus leading to a decrease in RBC count (Abdel-Tawwab et al. 2010). This is different from the result of Zhang (1991), who found that RBC count and haemoglobin concentration were increased as water temperature increased from 15 to 35 °C. Qiang et al. (2013) also found that WBC counts at high or low temperatures sideways, the optimal temperature range (29–31 °C) (Popma & lovshin 1995) was lower. Slater & Schreck (1998) suggested that acute temperature differences such as winter-summer temperature dynamics may inhibit immunological indices.

Morgan et al. (2008) found that RBC and WBC counts in blood of rainbow trout at a temperature range of 8–12 °C were higher than 0–3 °C temperature range. de Pedro et al. (2005) and Guijarro et al. (2003) found that RBC count and haemoglobin concentration of tench (Tinca tinca) were mainly affected by water temperature, and increased as water temperature rises; and hence showed that RBC count and haemoglobin concentration in summer were higher than in winter. Biochemical reactions in the fish body could have being quickened in an environment with optimum temperature range and increased oxygen supply, leading to the increase RBC products (Bowden et al. 2007). Meanwhile, DO would be more sufficient in low temperature water; the blood may become more viscous, and hence require less RBC to transport oxygen (Morgan et al. 2008). In other aquatic animals such as two scallop species, Argopecten irradians and Chlamys farreri, similar results were observed (Liu et al. 2004).

However, the change of haematological parameters in fish may be related to their tolerance. Some fish species have no sensitivity to temperature changes. For instance, when water temperature ranged over 15–31 °C, temperature had no influence on RBC count and haematocrit in silver catfish (Rhamdia quelen) (Lermen et al. 2004). Pascoli et al. (2011) also found that the WBC counts of sea bass and tench were significantly higher at suitable water temperature in summer than in winter.

The effect of cold temperature on blood parameters of Shizothorax richardsonii showed a decrease in RBC, WBC and haemoglobin content. This decreasing trend of haemoglobin in cold temperatures has also been reported in many species (Staurnes et al. 1994; Chen et al. 1995). This is because S. richardsonii survive at low temperature by concentration of Na+ and K+ content in plasma. Water temperature decrease, increase the osmotic pressure, increase Na+ and K+ content in plasma. Sodium and potassium levels increased during the initial decrease in water temperature in fish (Doudoroff 1945). Gill Na+, K+-ATPase activity is correlated to both active ion uptake and excretion (Evans 1982) and a general assumption is made that a significant increase in gill Na+, K+-ATPase activity is necessary to maintain normal electrolyte and water balance in seawater (McCormick et al. 1987).

Increased numbers of neutrophils in Atlantic salmon S. salar maintained at high temperature (18 °C) were also reported (Pettersen et al. 2005). In contrast, no alteration in RBC was found in Atlantic salmon exposed to 5 and 15 °C (Fivelstad et al. 2007). Lowering the water temperature from 23 to 11 °C in a period of 24 h resulted in anaemia but no alteration in neutrophil population in the blood of channel catfish (Bly & Clem 1991). However, decreased WBC counts have been found in hybrid striped bass Morone chrysops × Morone saxatilis maintained at high temperature (Hrubec et al. 1997) and in tilapia transferred from 27 to 19 °C and 27 to 35 °C (Ndong et al. 2007). When water temperature was 25 °C or higher, channel catfish showed increases in RBC, WBC, thrombocytes and monocytes (Martins et al. 2011a, b).

Physiological responses to chronic temperature changes are often such that they compensate, partially or completely, for the effects of temperature on the rate of body functions, that is, they tend to maintain metabolism, locomotion and cardiac function relatively independent of temperature changes (Tiitu & Vornanen 2001). Sometimes animals even actively exaggerate the effects of temperature by suppressing the rate of physiological processes more than would be caused by a simple temperature effect (Q10). This process is called inverse acclimation and usually leads to metabolic depression under harsh environmental conditions and thereby extends the survival time of the individual (Jackson 2000).

Temperature has a strong effect on the rate of all biological processes including the heart rate, rate of impulse conduction through the heart and shortening velocity of the cardiac muscle are strongly retarded by low temperatures and accelerated by high temperatures (Farrell & Jones 1992; Opie 1998). In both stenothermic (narrow thermal tolerance) and eurythermic (wide thermal tolerance) species the optimum temperature for heart function is close to the preferred body temperature of the fish, and the activity of the heart is depressed towards both low and high extremes of thermal tolerance (Brett 1971; Matikainen & Vornanen 1992). It is proposed that when the temperature approaches the upper or lower limits of thermal tolerance, oxygen delivery systems may fail to provide enough oxygen to the tissues and the animal especially fish will die of hypoxia or anoxia, mainly due to temperature-dependent limitation of the circulation.

Most fish are in thermal equilibrium with the surrounding water due to efficient counter current heat exchange between the water and blood at the gills (Stevens & Sutterlin 1976). Temperature is sensed in fish both in the brain and peripherally in the skin, and thermoregulatory behaviours are influenced by both (Crawshaw et al. 1985). The Bear Lake sculpin, Cottus extensus, by postprandially moving from colder (5 °C) to warmer water (15 °C), can triple their growth rate (Wurtsbaugh & Neverman 1988; Neverman & Wurtsbaugh 1994).

In the case of Paraphysa scrofa at 20 °C, regardless of the water pH in which the fish were kept, the increase of haematocrit and mean cell volume, with concomitant decreases in RBC, haemoglobin concentrations and mean cell haemoglobin concentration compared to the water pH 7.0 of controls, may indicate cell swelling (Carvalho & Fernandes 2006). In general, at a low temperature (20 °C), the change in water pH leads to haemodilution while, at a high temperature (30 °C), the changes in blood parameters demonstrate a response to improve the oxygen transport that should increase to maintain homeostasis (Carvalho & Fernandes 2006).

Fivelstad et al. (2007) found increased plasma pH (by about 0.1 pH units) with temperature but blood haematocrit and haemoglobin concentrations were not significantly different for the control groups kept at 5 and 15 °C. Data from short-term experiments on rainbow trout exposed to exhaustive exercise showed that temperature influenced the recovery pattern for plasma pH (Kieffer et al. 1994). Trout acclimated to 5 °C required more time for recovery than fish acclimated to 18 °C. Clearly, the physiological effects related to acid–base balance and high CO2 is different at different temperatures, most likely related to overall reduced energy metabolism at lower temperatures and changes in energy partitioning. The physiological impact of CO2 was lowest at the high temperature (Fivelstad et al. 2007).

Adam & Agab (2008) found the blood glucose, total plasma protein, uric acid and urea levels (76.74 ± 4.98, 4.08 ± 0.74, 1.57 ± 0.38 and 22.5 ± 3.45, respectively) of studied Clarias garipinus were similar to those reported by Tavares-Dias et al. (2000). However, these levels were significantly higher than the levels of channel catfish Ictalus metas (O'Neal & Weirich 2001). These variations might be attributed to seasonal temperature changes which may affect the biochemical blood levels, although the data refer to fishes from tropical regions where temperature was more or less stable throughout the year (Adeyemo et al. 2003). Recently, biomarkers of cellular stress have been widely used in the assessment of exposure of aquatic organisms to environment. Biomarkers of oxidative stress have been among the most commonly used biomarkers of cellular stress for habitat quality assessment, namely lipid peroxidation, catalase, superoxide dismutase and glutathione-S-transferase activities, used mainly as indicators of cellular stress resultant from environmental changes. However, environmental variables, strongly related to seasonality and extreme natural events, such as temperature and salinity, are known to have a significant effect on oxidative stress biomarkers (Amado et al. 2006; Vinagre et al. 2014). Roche & Bogé (1996) also tested the effect of temperature on oxidative stress biomarkers in Dicentrarchus labrax and concluded that lipid peroxidation and catalase activity were increased by thermal stress. They also investigated these biomarkers for chemical toxicity and concluded that although they respond to this stressor, they also respond to temperature and salinity variations.

EFFECTS OF TEMPERATURE ON GASTRIC EVACUATION RATE OF FISH

Ectothermic animals depend on external heat sources and appropriate behaviours to regulate body temperature in the face of environmental changes (Cowles & Bogert 1944). Many physiological functions and behaviours such as resting and maximal metabolic rates, locomotion and digestion are temperature dependent and fishes are constantly faced with the need to prioritize the selected body temperature to satisfy these different needs (Stevenson et al. 1985; Dorcas et al. 1997). Most animals must perform many tasks simultaneously, but since temperature sensitivity of different biological functions may vary, thermoregulation can impose potential conflicts between functions (Peterson et al. 1993). Lately, the scientific discussion, regarding the physiological effects of climate change on fish, has been focused on a hypothesis suggesting that the temperature limitations are mainly set by a reduction in metabolic scope (Claireaux & Lefrançois 2007; Portner et al. 2007; Wang & Overgaard 2007; Farrell 2009). Knowledge about food habits and total consumption is necessary for an understanding of the role of a fish species within the ecosystem. An energy budget approach alone does not provide information about the food habits. This can only come from a description of stomach contents of fishes sampled in the field. In order to convert the quantitative information on the stomach contents into estimates of ingestion rates, it is necessary to acquire knowledge about the rate at which the food items disappear from the stomachs (Andersen 1984). The term gastric evacuation rate (GER) denotes the rate at which food passes from the stomach into the intestine, even though this is something of a misnomer (Windell et al. 1978).

Gut evacuation rates have been mainly determined in fish during either field or laboratory experiments (Jobling 1980a, b; Heroux & Magnan 1996). There are many factors which have an impact on fish gastric evacuation (GE). Of them, the most significant ones are temperature, fish size, feed type, feed amount and feed quality (Andersen 1998; Koed 2001). According to Elliott (1972) and Irigoien (1998), the initial gut content should not significantly affect the evacuation rate, with temperature being the limiting factor. A combination of stomach evacuation rates with temperature and field estimates of stomach fullness is commonly used to estimate daily rations of fish in the wild (Sweka et al. 2004; Kawaguchi et al. 2007).

Some studies indicate instantaneous evacuation rates (re) increase exponentially with temperature (Elliott 1972; Persson 1981; From & Rasmussen 1984), whereas other studies indicate that evacuation rates increase linearly with temperature (Ryer & Boehlert 1983; Worobec 1984). An exponential increase in instantaneous evacuation rate with temperature indicates that the range in food weight passed through the pylorus will be greater than if temperature affects evacuation rate linearly.

In squawfish, GER increased from 5% of stomach volume/hr at 4–6 °C to 40–50%/hr at 24 °C (Steigenberger & Larkin 1974). In rainbow trout, the greatest absolute increases were at the low end of the temperature range rather than at the high end. Depending on food type, it took three to four times as long to empty the stomach by either 50 or 100% at 5 °C than at 20 °C (Windell et al. 1976). For channel catfish the fastest GER occurred at 26.6 °C, with slower rates at lower and higher (29.4 °C) temperatures (Shrable et al. 1969). The authors suggested that their force-feeding method might have been stressful and the stress would have been most noticeable at higher temperatures.

Brett (1971) presented an interesting hypothesis concerning temperature effects on digestion efficiency to explain a diurnal migration made by sockeye salmon juveniles in Lake Babine (British Columbia, Canada) during the summer. When this oligotrophic lake has strong thermocline, juvenile sockeye feed at the surface in warm (12–15 °C) water at dawn and dusk, but descend into cooler (4–6 °C) water to digest their meal. Brett had shown earlier that sockeye salmon juveniles have increased digestion (assimilation) efficiency at lower temperatures and lower rations and thus hypothesized that they were maximizing their assimilation efficiency and growth by making the vertical migration. Biette & Geen (1980) tested the hypothesis by growing underyearling sockeye salmon at 5.5, 15.8 °C and a cyclic combination of the two temperatures. Fish grew more rapidly under the cyclic than under either constant temperature when eating 14C-labelled Daphnia. This suggests that Brett's hypothesis concerning behavioural temperature control of their digestive processes was correct.

The predictable scaling of fish metabolic rate with temperature and ontogeny is believed to be the primary driver of changes in rates of feeding and gastric evacuation (Paloheimo & Dickie 1966). Roughly two-fold increases in maximum consumption rate were observed as water temperature varied between 17 and 27 °C for both small and large body sizes of red drum. Similarly, Hartman & Brandt (1995) demonstrated one- to four-fold increases in maximum consumption rates between approximately 5 and 30 °C for three temperate fish predators in Chesapeake Bay, USA, with two- to three-fold variation in consumption across body sizes that ranged from roughly 10 to 1,000 g.

The effects of water temperature were strongest with red drum achieving faster rates of evacuation when held at the higher temperature tested. The influence of water temperature on fish metabolism and rates of meal processing has been studied extensively (Buckel & Conover 1996; Wuenschel & Werner 2004), with a general increase, often of exponential form, observed in gut evacuation rate at higher temperatures.

Temperature had a strong effect on gut evacuation rates of brown trout and digestion rate increased 2.1 times for each 10 °C increase in temperature (the Q10) (He & Wurtsbaugh 1993). Elliott (1972, 1991) found that increasing temperature had a much stronger effect on digestion rates of brown trout feeding on invertebrates (Q10 = 3.2) or fish fry (Q10 = 3.0) than He & Wurtsbaugh (1993) found (Q10 = 2.1). At temperatures above 10 °C, this results in 20–40% differences in estimated digestion rates between the different studies. Durbin et al. (1983) reviewed individual temperature coefficients for several marine and freshwater fish and found a mean temperature coefficient of 0.115, which also gives a very high Q10 of 3.2. Fange & Grove (1979) reported a Q10 of 2.6 in their analysis of digestion in fishes. The reasons for the substantial difference between He & Wurtsbaugh (1993) and those of these researchers is not clear.

In roach (Rutilus rutilus), increasing water temperature resulted in heightening enzyme activity and feed intake rates (Hardewig & van Djik 2003). Cooler water temperatures may reduce digestion rates and increase gut transit time (lowering gastrointestinal evacuation rates), thus nutrient digestibility will suffer (Miegel et al. 2010). Protease enzymes in the gut are not as active at lower temperatures (Kofuji et al. 2005). Trypsin activity levels are lower during the winter months compared to spring months in Atlantic salmon (Einarsson et al. 2005). Furthermore, Alexander et al. (2011) showed that higher water temperatures resulted in higher amylase, protease, and hexokinase activities in rohi (Labeo rohita) fingerlings. However, McLeese & Stevens (1982) showed that temperature did not have a significant effect on trypsin and chymotrypsin activity in rainbow trout.

The rate of gastric emptying has been positively related with the return of appetite and food consumption in fish species such as rainbow trout, O. mykiss (Grove et al. 1978), winter flounder, Pseudopleuronectes americanus (Huebner & Langton 1982), rockfish, Sebastes schlegeli (Lee et al. 2000), dab, Limanda limanda (Gwyther & Grove 1981) and turbot, S. maximus (Grove et al. 1985).

Temperature had a strong negative effect on gastric evacuation in both Atlantic cod and haddock (Perez-Casanova et al. 2009). This was not surprising as dos Santos & Jobling (1995) showed that the Q10 for Atlantic cod gastric emptying was 3.7 and studies on numerous other fish species have shown GE to be highly temperature dependent: brown trout, Q10 = 3.0 (Elliott 1972); whiting, Q10 = 2.2 (Andersen 1999); plaice, Q10 = 2 (Basimi & Grove 1985). It is very likely that decrease in GE between 11 and 2 °C was due to both the direct effect of temperature on GE, and the large reduction in the amount of food consumed between the two temperatures (dos Santos & Jobling 1991).

Nakada (2000) reported that gut transit time in Japanese yellowtail significantly increased when water temperatures and feeding frequency decreased while conversely Kofuji et al. (2005) reported a decreased digestion rate in the same species during the colder months. They also described decreased trypsin and chymotrypsin secretion in colder water temperatures resulting in lower apparent protein digestibility. This was supported in a study by Satoh et al. (2004) which revealed that apparent protein digestibility in Japanese yellowtail was significantly lower in seasons when water temperatures were colder. Although enzyme activity in the stomach of Japanese yellowtail was higher in warmer water temperatures, intestinal enzyme activity was greater in colder water temperatures, possibly due to slower movement of digesta through the intestines during the winter months (Kofuji et al. 2005).

It took approximately three times as long for all digesta to be voided from the fish in winter compared to summer (Miegel et al. 2010). This is consistent with previous studies with Japanese yellowtail (Nakada 2000; Watanabe et al. 2001), in which water temperature has been shown to influence gut transit time and digestion rates.

CONCLUSION

Current data show global climate change effects ranging from changes in metabolic rates, growth, physiology and survival of the fish community. However, the implications for performance, development and plasticity are only recently being realized. Response to environmental changes must be considered within the context of an organism's normal environment, as thermal sensitivity is generally a reflection of field temperature and levels of local adaptation. To date, many of the predictions regarding the biology and ecology of marine fish come from models of temperate systems and research has been carried out indiscriminately in captive conditions especially in temperate regions; but, in contrast, for tropical fish species very little research has been conducted. Our results imply that temperature changes manifest themselves in a variety of ways in different life stages and in different conditions and the different magnitude of response in growth, physiology, development and survival emphasizes the value of using measurements appropriate to the ecology of the organism. Marine and ocean ecosystems provide more adaptable potential for fish in the event of climate change (e.g., ocean upwelling, tides and deep-ocean circulation transform heat and cold water between low and high latitudes) resulting in stabilizing temperature and nutrient transport. The increased surface temperature will proportionally increase change in water temperature, and thus the salinity and limnology of the water. On the other hand, closed systems such as lakes or lagoons can be less adaptable because of abrupt changes in water parameters during storm surge or cyclone. With a broader and deeper information base, researchers will be able to more accurately predict future situations. Without an understanding of how these organisms and systems function and interact, we cannot predict how they will react to perturbation, including global climate change-related disturbances. These gaps lead to uncertainties about future fish stocks and for the people depending on them.

ACKNOWLEDGEMENTS

We would like to thank the two anonymous reviewers for their very useful comments that greatly improved the quality of the manuscript. We also would like to thank Universiti Kebangsaan Malaysia (UKM) for the Research University ZAMALAH scholarship provided to S. K. Mazumder and Moumita De. This study was supported by UKM through the research grant # GUP-2013-006, Ministry of Science Technology and Innovation, Malaysia through the e-Science grant # 04-01-02-SF1208, and Ministry of Higher Education, Malaysia through the Exploratory research grant # ERGS/1/2013/STG03/UKM/01/2.

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