**Fatty Acids' Profiles of Aquatic Organisms: Revealing the Impacts of Environmental and Anthropogenic Stressors**

Ana M.M. Gonçalves, João C. Marques and

Fernando Gonçalves

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/intechopen.68544

#### **Abstract**

There is a great concern about the impacts of climate changes namely due to salinity sea‐ water and temperature alterations in aquatic organisms with the estuarine and coastal environments being the major affected areas. The intensive usage of chemicals in an indiscriminate way in agriculture practices, achieving, in some cases, values above the limits of contamination authorized by the European legislation, also drastically affects the surrounded estuarine areas with profound consequences to the water quality and the aquatic communities. It is known that stressors affect organisms' physiological conditions with recent works concerning alterations in the fatty acid (FA) profiles associated with environmental and contamination events that become more frequent. FA plays a key role in immune and physiological functions and is associated with the prevention of some diseases, shown to be good bio‐indicators to assess the organisms' impacts under stress conditions. Thus, this chapter proposes to address natural (salinity and temperature) and chemical (herbicide and metal) stressors' impacts in the FA profiles of *Thalassiosira weissflogii* and *Cerastoderma edule* and infers about the effects on organisms' physiological processes and along the food web. Consequences in food resources and to healthier and nutritious food consumption with benefits to human beings are also assessed.

**Keywords:** fatty acids, bio‐indicator, stressors, climate changes, salinity, temperature, pollutants, herbicide, metal, aquatic organisms, food quality, estuaries

© 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

#### **1. Introduction**

Fatty acids (FAs) are essential molecules with a crucial role in the maintenance of physiologi‐ cal functions of many organisms. These carboxylic acids provide fuel for the brain and the tissues at the metabolic level and are a major constituent in the cellular wall as part of phos‐ pholipids. Once transferred across the food chain, FAs perform the connection between pri‐ mary producers and secondary consumers [1]. Greatly abundant in the brain tissues, these molecules represent almost half of the brain weight [2]. Fatty acid nomenclature is represented by X:YωZ, where X represents the number of carbon atoms in the chain, Y represents the num‐ ber of double bonds and Z gives information about the position of the first double bond count‐ ing from the methyl group [3]. There are two major groups of FA: saturated fatty acids (SFAs) and unsaturated fatty acids (UFAs), differing between them by the existence of double bonds in UFA. The position of the double bonds is determined by the desaturase enzyme activity, which performs the double bond in different positions accordingly to the type of enzyme pres‐ ent in different organisms (e.g. Δ3 in animals, Δ6 in animals and plants and Δ15 in algae and plants with chlorophyll) [3]. Saturated FAs are metabolized mainly as a source of energy and also as lipid storage. UFA can be identified by the number of double bonds: Monounsaturated FA (MUFA) have one double bond and can be synthesized *de novo* almost by all organisms [4]. Polyunsaturated FAs (PUFAs) are fatty acids with two or more double bonds that play an essen‐ tial role in the brain development [5] and many physiological functions such as down regulat‐ ing inflammation, cellular signalling [6] and regulation of transcription factors [7]. Essential fatty acids (EFAs) are some PUFAs that play major important functions in physiological and biochemical processes and that must be acquired externally, through dietary input, once the majority of the animals cannot synthesize them *de novo* [8], due to the lack of the desaturase enzyme [9]. Although some animals are able to synthetize EFA from linolenic precursors such as α‐linolenic acid C18:3 (n‐3), C18:4 (n‐3) and C18:5 (n‐3), found almost exclusively in plants [3] via elongation and desaturation, the rate this conversion succeeds is residual to supply the necessary amount of EFA required for an optimal growth and development. Thus, EFAs must be obtained by direct feeding on phytoplanktonic, plants or bacteria species [10], or by ingesting lipid emulsions with high content of EFAs [11]. Since EFA are later transferred along the food web, aquatic species like fish and other organisms from higher trophic levels are an important food sources of such molecules [4]. EFAs are represented mainly by some PUFAs. Highly unsaturated fatty acids (HUFAs) are a subset of PUFAs with a chain of 20 carbon atoms or more and with three or more double bonds that play an important role in cellular growth, with special relevance in tissue growth, energy storage, neural development and also repro‐ ductive fitness [4, 12]. The physiological activities of EFAs in animals are mainly represented by eicosapentaenoic acid (EPA—20:5n3) and docosahexaenoic acid (DHA—22:6n3), two of the most important HUFAs that are synthesized *de novo* by phytoplankton and bio‐accumulated by animals. The arachidonic acid (ARA—20:4n‐6) is also a representative EFA with functions as a precursor of animal hormones such as prostaglandins and leukotrienes amongst others [3]. EPA and DHA play a major role in brain development and maintenance of brain structure and function [13]. EPA intake influences many physiological processes such as reproduction, immunity efficiency and osmoregulation [3]. DHA is important for the health and developing of neurons and for neurotransmission, with strong influence on cognition and behavior, and it is also proved to be important in the protection against oxidative stress [14]. Fatty acids are considered to be an accurate tool in trophic interaction studies [15], mainly due to their impor‐ tance in the health/stability of the ecosystem, and because they are transferred conservatively to higher trophic levels along the trophic food web [4, 16]. Furthermore, FA profiles can reflect structural changes in species' biochemical composition in response to stressors [17–20]. Lipid components are also very sensitive to environmental changes, which make them an efficient assessment tool to monitor toxicological effects on the marine biota as bio‐indicators of eco‐ system health [19].

**1. Introduction**

90 Fatty Acids

Fatty acids (FAs) are essential molecules with a crucial role in the maintenance of physiologi‐ cal functions of many organisms. These carboxylic acids provide fuel for the brain and the tissues at the metabolic level and are a major constituent in the cellular wall as part of phos‐ pholipids. Once transferred across the food chain, FAs perform the connection between pri‐ mary producers and secondary consumers [1]. Greatly abundant in the brain tissues, these molecules represent almost half of the brain weight [2]. Fatty acid nomenclature is represented by X:YωZ, where X represents the number of carbon atoms in the chain, Y represents the num‐ ber of double bonds and Z gives information about the position of the first double bond count‐ ing from the methyl group [3]. There are two major groups of FA: saturated fatty acids (SFAs) and unsaturated fatty acids (UFAs), differing between them by the existence of double bonds in UFA. The position of the double bonds is determined by the desaturase enzyme activity, which performs the double bond in different positions accordingly to the type of enzyme pres‐ ent in different organisms (e.g. Δ3 in animals, Δ6 in animals and plants and Δ15 in algae and plants with chlorophyll) [3]. Saturated FAs are metabolized mainly as a source of energy and also as lipid storage. UFA can be identified by the number of double bonds: Monounsaturated FA (MUFA) have one double bond and can be synthesized *de novo* almost by all organisms [4]. Polyunsaturated FAs (PUFAs) are fatty acids with two or more double bonds that play an essen‐ tial role in the brain development [5] and many physiological functions such as down regulat‐ ing inflammation, cellular signalling [6] and regulation of transcription factors [7]. Essential fatty acids (EFAs) are some PUFAs that play major important functions in physiological and biochemical processes and that must be acquired externally, through dietary input, once the majority of the animals cannot synthesize them *de novo* [8], due to the lack of the desaturase enzyme [9]. Although some animals are able to synthetize EFA from linolenic precursors such as α‐linolenic acid C18:3 (n‐3), C18:4 (n‐3) and C18:5 (n‐3), found almost exclusively in plants [3] via elongation and desaturation, the rate this conversion succeeds is residual to supply the necessary amount of EFA required for an optimal growth and development. Thus, EFAs must be obtained by direct feeding on phytoplanktonic, plants or bacteria species [10], or by ingesting lipid emulsions with high content of EFAs [11]. Since EFA are later transferred along the food web, aquatic species like fish and other organisms from higher trophic levels are an important food sources of such molecules [4]. EFAs are represented mainly by some PUFAs. Highly unsaturated fatty acids (HUFAs) are a subset of PUFAs with a chain of 20 carbon atoms or more and with three or more double bonds that play an important role in cellular growth, with special relevance in tissue growth, energy storage, neural development and also repro‐ ductive fitness [4, 12]. The physiological activities of EFAs in animals are mainly represented by eicosapentaenoic acid (EPA—20:5n3) and docosahexaenoic acid (DHA—22:6n3), two of the most important HUFAs that are synthesized *de novo* by phytoplankton and bio‐accumulated by animals. The arachidonic acid (ARA—20:4n‐6) is also a representative EFA with functions as a precursor of animal hormones such as prostaglandins and leukotrienes amongst others [3]. EPA and DHA play a major role in brain development and maintenance of brain structure and function [13]. EPA intake influences many physiological processes such as reproduction, immunity efficiency and osmoregulation [3]. DHA is important for the health and developing

Environmental pollution worldwide is an undesirable by‐product of the increased demand for natural resources in the modern civilization. However, since the advent of human societ‐ ies, there have always been foci of environmental contamination, though nothing on the scale we see today. Practically, whole environment suffers from some degree of contamination in concentrations above those expected for the region. The pollutants that damage the ecosys‐ tem are the pollutants from industry and mining that release toxic substances such as metals and organic pollutants. Some pesticides and mainly metals (e.g. Cd, Cr, Pb, Hg, Ni, Cu) are non‐degradable and therefore accumulate in nature, where they continue to affect ecosys‐ tem's function over the course of decades or even centuries. These chemicals can distress several biological organization levels affecting flora and fauna aquatic organisms, interfering with the metabolic and physiological processes and thus compromising the structure and physicochemical properties of the membrane, damage cells, tissues and organs. Long‐term effects may lead to higher mortality among population, changing the diversity and structure of the communities. Furthermore, due to global climate changes, environmental conditions are expected to change considerably in several areas. In some regions, it is expected, seasonal differences become more notorious than they used to be exposing the organisms to a wide physiological stress. These changes not only act as additional stress factors but may also con‐ siderably modify the toxicity of pollutants in aquatic ecosystems.

Estuaries are coastal ecosystems, which are biologically highly productive and having great importance in the ecological and at socioeconomic contexts, providing exceptional natural resources and services to human beings, mainly to local populations. Some of these sys‐ tems are located near farmlands, industrial and residential areas being under anthropogenic pressures that affect water quality and the aquatic communities. Estuarine systems are very useful model systems to study the ecological and evolutionary responses of organisms to highly variable, discontinuous habitats due to the extreme daily variations that occur in these transitional areas exposing the organisms to a widely physiological stress [21]. The transition between the freshwater and marine environments creates a gradient of physi‐ cal and chemical conditions that determine the amount and distribution of the species and communities that live at these ecosystems [22], with salinity being one of the major control‐ ling factor of species' distribution in estuarine systems. Aquatic organisms from these eco‐ systems are exposed to physical and chemical environmental conditions that vary greatly, on both seasonally and daily basis. Because planktonic species are strongly influenced by climatic factors, and particularly sensitive to changes in hydrological conditions [23], the increase in frequency of flooding episodes has a strong impact on macrobenthic communi‐ ties and is proven to be result into a significant decline in the diversity of suspension‐feeder species, such as microcrustaceans (e.g. copepods) [24]. The temperature of the water is also linked with changes in salinity, and although some studies have tested the two parameters separately, both should also be studied in a bi‐dimensional approach, to best simulate natu‐ ral conditions [25]. According to Kinne [26] and Williams and Geddes [27], temperature can alter the physiological tolerance of an organism to salinity changes, and in turn, salinity can influence the impact of environmental temperature on the same organism. Salinity is indeed of major importance in the distribution of aquatic organisms because the ability of osmo‐ regulation affects ecological tolerances and the type of ecosystem (marine, coastal, estuarine, freshwater) [28].

Since past decades, extreme weather episodes are frequent worldwide, and Portugal is not an exception. At the Mondego estuary, located in the western coast of Portugal, near Figueira da Foz city, episodes of drought and flood have been registered and are well documented in the literature revealing ecological impacts on aquatic communities [24, 29–31]. This temperate estuarine system surrounded by agriculture fields, the commercial port, beach and industries, and with a high marine exploitation of resources, suffers high anthropogenic pressures similar to many other estuarine systems mainly from the Mediterranean region. Rice and corn fields are the main agriculture production in the Mondego valley, being Viper and Primextra® Gold TZ the most used pesticides in agriculture practices, respectively, according to information from the cooperatives of the region. Furthermore, copper is one of the main constituents of pesticides formulations, with application in agricultural activities. It is an essential metal, with vital importance in low concentrations to organisms, acting as a co‐factor of many enzymes, i.e. it is a component of superoxide dismutase, an enzyme defending living organisms against reactive oxygen species [32]. Still becomes toxic at high concentrations affecting several bio‐ chemical and metabolic processes such as FA metabolism, cell division, photosynthesis, respiration and synthesis of carbohydrates, pigments and chlorophyll [33, 34]. In 1998, and similarly to other estuarine systems near intensive agriculture practices with wide usage of pollutants, a pesticide‐monitoring program was implemented in Mondego estuary to recover the system [35].

Stressors affect organisms' growth and biochemical processes and also their performance and healthy status. To compensate extreme conditions, or at least conditions that are far from the optimal, some organisms developed strategic and adaptive mechanisms to com‐ pensate physiological requirements. Nevertheless, it is apparent to have the occurrence of significant losses and large alterations on the FA contents. Some studies allow to rec‐ ognize and assess the response of specific markers in order to identify and validate pre‐ cise bio‐indicators that are able to capture the impact of disturbances resulted at extreme conditions or at the presence of stressors, which might be used as early warning signals of stressing conditions and, eventually, be associated to strategic prevention of stress‐asso‐ ciated diseases. Therefore, it is crucial to determine and assess lethal effects and physio‐ logical responses of aquatic organisms under the influence of environmental (e.g. salinity, temperature) and chemical (e.g. pollutants) stressors in order to predict the impacts on communities and thus on aquatic ecosystems and food quality. Still, there are some char‐ acteristics that must be taken into account when choosing the species to be tested, such as (1) the species' sensitiveness to the studied parameter/substance; (2) well‐known nutrient requirements; (3) low genetic and phenotypic variability among strains/organisms; and (4) fast and cost efficient maintenance in the laboratory [36]. The marine phytoplankton species, *Thalassiosira weissflogii*, is often used as a bio‐indicator in several studies, becom‐ ing increasingly relevant in environmental monitoring studies [36]. The marine diatom value is closely related to its many applications in 1) ecotoxicological studies; 2) biodiesel production; 3) prey for zooplankton (rotifers, copepods, brine shrimp) and 4) produc‐ tion of metabolites such as lipids, proteins, carbohydrates, pigments and vitamins [37]. *Cerastoderma edule* plays a key role between primary producers and consumers. They live in intertidal shallow areas, presenting a suspension‐feeder behavior [38]. This bivalve spe‐ cies lives worldwide occurring from the Northern Norway to the North Africa, on the east coast of the Atlantic and in Murmansk in the Arctic [39]. Due to its sessile life style, easy sampling collection, maintenance, handling and sensitivity to chemicals, *C*. *edule* is widely used as standard species in ecotoxicological bioassays [19, 20]. Due to its high ability to filtrate and accumulate large amount of pollutants, this species is also used as bio‐indica‐ tor in ecological studies [39–43]. Furthermore, *C. edule* is very much appreciated as food source mainly by local populations, which highlights its importance to the socioeconomic sector.

In this chapter, it is proposed to determine and assess the effects of environmental (salinity and temperature) and chemical (Primextra® Gold TZ and Copper) stressors, individually and combined, in the fatty acid profiles of a marine phytoplankton species (*T. weissflogii*) and an estuarine bivalve species (*C. edule*). The impacts of global stressors to the quality of aquatic food resources and thus to a healthy and nutritive food consumption are also assessed.

### **2. Material and methods**

increase in frequency of flooding episodes has a strong impact on macrobenthic communi‐ ties and is proven to be result into a significant decline in the diversity of suspension‐feeder species, such as microcrustaceans (e.g. copepods) [24]. The temperature of the water is also linked with changes in salinity, and although some studies have tested the two parameters separately, both should also be studied in a bi‐dimensional approach, to best simulate natu‐ ral conditions [25]. According to Kinne [26] and Williams and Geddes [27], temperature can alter the physiological tolerance of an organism to salinity changes, and in turn, salinity can influence the impact of environmental temperature on the same organism. Salinity is indeed of major importance in the distribution of aquatic organisms because the ability of osmo‐ regulation affects ecological tolerances and the type of ecosystem (marine, coastal, estuarine,

Since past decades, extreme weather episodes are frequent worldwide, and Portugal is not an exception. At the Mondego estuary, located in the western coast of Portugal, near Figueira da Foz city, episodes of drought and flood have been registered and are well documented in the literature revealing ecological impacts on aquatic communities [24, 29–31]. This temperate estuarine system surrounded by agriculture fields, the commercial port, beach and industries, and with a high marine exploitation of resources, suffers high anthropogenic pressures similar to many other estuarine systems mainly from the Mediterranean region. Rice and corn fields are the main agriculture production in the Mondego valley, being Viper and Primextra® Gold TZ the most used pesticides in agriculture practices, respectively, according to information from the cooperatives of the region. Furthermore, copper is one of the main constituents of pesticides formulations, with application in agricultural activities. It is an essential metal, with vital importance in low concentrations to organisms, acting as a co‐factor of many enzymes, i.e. it is a component of superoxide dismutase, an enzyme defending living organisms against reactive oxygen species [32]. Still becomes toxic at high concentrations affecting several bio‐ chemical and metabolic processes such as FA metabolism, cell division, photosynthesis, respiration and synthesis of carbohydrates, pigments and chlorophyll [33, 34]. In 1998, and similarly to other estuarine systems near intensive agriculture practices with wide usage of pollutants, a pesticide‐monitoring program was implemented in Mondego estuary to recover

Stressors affect organisms' growth and biochemical processes and also their performance and healthy status. To compensate extreme conditions, or at least conditions that are far from the optimal, some organisms developed strategic and adaptive mechanisms to com‐ pensate physiological requirements. Nevertheless, it is apparent to have the occurrence of significant losses and large alterations on the FA contents. Some studies allow to rec‐ ognize and assess the response of specific markers in order to identify and validate pre‐ cise bio‐indicators that are able to capture the impact of disturbances resulted at extreme conditions or at the presence of stressors, which might be used as early warning signals of stressing conditions and, eventually, be associated to strategic prevention of stress‐asso‐ ciated diseases. Therefore, it is crucial to determine and assess lethal effects and physio‐ logical responses of aquatic organisms under the influence of environmental (e.g. salinity, temperature) and chemical (e.g. pollutants) stressors in order to predict the impacts on

freshwater) [28].

92 Fatty Acids

the system [35].

#### **2.1. Study area and sampling procedure**

The Mondego estuary is a small mesotidal system in the West Atlantic coast of Portugal (40°08′N, 8°50′W). The estuary is divided into two arms, north and south (**Figure 1**). The northern arm is characterized by a salt‐wedge during low tide, which changes to a par‐ tially mixed water column at high tide. It is characterized by a partially mixed water col‐ umn at low tide and a well‐mixed one at high tide at spring tides [44]. The southern arm is shallower and its water circulation is mostly dependent on tides and on freshwater input from a small tributary system, the Pranto River. Freshwater discharges of this river are controlled by a sluice according to the water needs by the rice fields of Mondego valley [29].

*C. edule* was sampled at the south arm of the estuary (**Figure 1**). Organisms were transported from the field in cold boxes with brackish water.

**Figure 1.** The Mondego estuary location and the sampling site within the estuary.

#### **2.2. Laboratory procedures and bioassays**

#### *2.2.1. Microalga species*

#### *2.2.1.1. Culture maintenance and laboratory bioassays*

*T. weissflogii* was obtain from the Scottish Marine Institute, Dunbeg, PA37 1QA, UK (strain number 1085/18). The microalga was maintained under laboratory conditions by renewing the alga medium once a week, maintaining a cell density of 2 × 10<sup>4</sup> cells/mL. f/2 medium was prepared accordingly to Guillard and Reyther [45] with water collected from the Mondego estuary with a salinity of 30 psu, previously filtered with Whatman glass microfiber filters with 1.2 μm pores and stored at 4°C. By the experiments with copper, the medium was prepared without EDTA, adapted after Rippingale and Payne [46]. A renew of algae culture was done weekly. All assays and organism cultures were maintained under artificial light with photo‐ period of 16hL:8hD. Different cultures were maintained in 15, 20 and 25 ± 2°C, respectively.

Before the beginning of bioassays, an inoculum of *T. weissflogii* was harvested from the bulk culture and incubated for 3 days in a chamber with photoperiod (16hL: 8hD) at 20°C [18]. This procedure was repeated from other two temperatures (15 and 25°C) to the salinity experi‐ ments where a set of different temperatures (15, 20 and 25°C) were assessed to a range of salinity concentrations. The cellular concentration was then adjusted to 10<sup>4</sup> cells/mL after cellular density determination at a Neubauer haemocytometer, and the microalgae was then exposed to a range of salt and chemical concentrations, respectively. Eight salinity concentra‐ tions (0, 5, 10, 15, 20, 25, 33) plus the control were performed at three distinct temperatures (15, 20 and 25°C). The experiments with the both toxicants were conducted to a range of concentrations from 0.200 to 0.800 mg/L of copper(II) sulphate pentahydrate and 0.005 to 0.040 mg/L of the herbicide Primextra® Gold TZ, plus the control. Glass beakers were used to saline and herbicide experiments, whereas plastic beakers were used in the experiments with the metal. Three replicas per treatment were prepared in each bioassay. The experi‐ ments were conducted under photoperiod (16hL: 8hD), with a duration of 96 h. At the end of the experiment, the cellular density was determined to each treatment using a Neubauer haemocytometer.

#### *2.2.1.2. Microcosm bioassays*

Microcosm bioassays had a duration of 7 days and has been conducted under the same con‐ ditions of the laboratory bioassays previously described. Three treatments corresponding to 96h‐EC10, EC20 and EC50 calculated from the bioassays described in the subsection above plus the control treatment were employed whenever possible due to the duration of the microcosm experiments. According to the results of the microalga growth obtained at the previous exper‐ imental bioassays, the control treatments of the salinity microcosm bioassays were performed to a salinity concentration of 33, 33 and 25 to the temperature of 15, 20 and 25°C, respectively. Although at 20°C the microalga presented a similar growth at the saline treatments of 30 (CTL) and 33, and being considered the optimal growth to *T. weissflogii* at the salinity of 30, the control treatment to the microcosm bioassay was performed to a salinity concentration of 33 in order to compare the results with the microcosm experiments conducted at 15°C. Three replicas per treatment were conducted. At the end of the microcosm bioassays, a final concen‐ tration of 7.2 × 106 cells/mL was measured in each Erlenmeyer flask and then filtered through a GF/F Whatman filter and frozen at −80°C for FA analysis.

#### *2.2.2. Bivalve species*

**2.2. Laboratory procedures and bioassays**

*2.2.1.1. Culture maintenance and laboratory bioassays*

the alga medium once a week, maintaining a cell density of 2 × 10<sup>4</sup>

**Figure 1.** The Mondego estuary location and the sampling site within the estuary.

*T. weissflogii* was obtain from the Scottish Marine Institute, Dunbeg, PA37 1QA, UK (strain number 1085/18). The microalga was maintained under laboratory conditions by renewing

prepared accordingly to Guillard and Reyther [45] with water collected from the Mondego estuary with a salinity of 30 psu, previously filtered with Whatman glass microfiber filters with 1.2 μm pores and stored at 4°C. By the experiments with copper, the medium was prepared without EDTA, adapted after Rippingale and Payne [46]. A renew of algae culture was done weekly. All assays and organism cultures were maintained under artificial light with photo‐ period of 16hL:8hD. Different cultures were maintained in 15, 20 and 25 ± 2°C, respectively.

Before the beginning of bioassays, an inoculum of *T. weissflogii* was harvested from the bulk culture and incubated for 3 days in a chamber with photoperiod (16hL: 8hD) at 20°C [18]. This procedure was repeated from other two temperatures (15 and 25°C) to the salinity experi‐ ments where a set of different temperatures (15, 20 and 25°C) were assessed to a range of

cellular density determination at a Neubauer haemocytometer, and the microalgae was then exposed to a range of salt and chemical concentrations, respectively. Eight salinity concentra‐ tions (0, 5, 10, 15, 20, 25, 33) plus the control were performed at three distinct temperatures (15, 20 and 25°C). The experiments with the both toxicants were conducted to a range of concentrations from 0.200 to 0.800 mg/L of copper(II) sulphate pentahydrate and 0.005 to 0.040 mg/L of the herbicide Primextra® Gold TZ, plus the control. Glass beakers were used

salinity concentrations. The cellular concentration was then adjusted to 10<sup>4</sup>

cells/mL. f/2 medium was

cells/mL after

*2.2.1. Microalga species*

94 Fatty Acids

#### *2.2.2.1. Culture maintenance and laboratory bioassays*

In the lab, organisms were divided in aquaria with aeration and filtrated sea water at the salinity of 20. On 10 selected organisms, collected in the field and not under any laboratorial process, a set of measurements (shell length, total weight, tissue weight and foot weight) were assessed to determine the condition indices. After the measurements, the muscle (foot) of each organism was removed and stored at −80°C for fatty acid analysis. The remaining organ‐ isms collected in the field were maintained in the aquaria, under photoperiod conditions (12hL:12hD) and control temperature (20 ± 2°C), without food, during a depuration period of 48 h, previously to the experiments.

Salinity bioassay was performed on organisms exposed to a range of saline concentrations from 0 to 35 plus the control. The salinity concentrations were obtained from successive dilutions of filtrated seawater at the salinity of 35 in distilled water. Experiments with the contaminants were conducted on individuals under six concentrations of copper(II) sulphate pentahydrate ranging from 0.6 to 2.1 mg/L and a set of eight concentrations from 0.5 to 60 mg/L of the herbi‐ cide Primextra® Gold TZ plus the control, respectively. The test medium was used as negative control. Bioassays were conducted under control temperature (20 ± 2°C), 12hL:12h<sup>D</sup>photoperiod, with filtrated sea water medium at the salinity of 20, during 120 h. Tests were carried out in glass (to saline and herbicide experiments) and plastic (to the metal) vials, 10 per treatment, containing a final test volume of 1000 mL per replicate. Organisms were fed daily with a com‐ mercial mixture of microalgae and rotifers. Organisms were transferred to newly prepared test solutions every next day. Bivalves were checked daily for mortality and behavioural conditions (to evaluate the conditions of the valves, organism behavior during feeding and the activity of the siphon). After the exposure period, all survival organisms were dissected, measured the weight and the body length and evaluated the condition indices of each individual. After the measurements, the muscle tissue (foot) was stored at −80°C for further fatty acid analysis.

#### *2.2.3. Fatty acid analysis*

The sample extraction for the FA analysis was obtained accordingly to the method described in Gonçalves et al. [19], substituting BF3‐methanol by H<sup>2</sup> S0<sup>4</sup> , due to reported deficiency in PUFA detection [47]. A differentiated phase was extracted, and an internal standard (fatty acid Methylnonadecanoate (C19) Fluka 74208) was added to the quantification of FA. The samples were later analyzed using a gas chromatograph with a mass spectrometer and an HP88 column (60 m × 25 mm × 0.20 μm). It was conducted in splitless mode, with a 1 μL injector per run. The column temperature was set to increase from 75 to 230°C at a rate of 2°C/min. The carrier gas was helium at a flow rate of 1.3 mL/min. The results of the GC analyses were obtained, and fatty acid methyl esters (FAMEs) were identified by compari‐ son of their retention times with those of individual purified standards. FAMEs can also be quantified by determining the area of the peaks of each fatty acid with the help of calibration factors [47].

#### *2.2.4. Statistical analysis*

The cellular density of *T. weissflogii* measured with a Neubauer Haemocytometer, at the end of the bioassays, was used to estimate the concentrations which induced *x*% growth inhibition (EC*<sup>x</sup>* values, with *x* = 10, 20, 50) and the corresponding 95% confidence intervals by non‐linear regression, using the least‐squares method to fit the data to the logistic equation.

The LC10, LC20 and LC50 values with corresponding 95% confidence intervals for *C. edule* were determined using Probit analysis [48].

To determine significant differences between treatments, one‐way analysis of variance (ANOVA) was performed, followed by Dunnett's multiple comparison test to identify signifi‐ cant differences between salinity treatments and the control treatment, considering a level of significance of 0.05.

The FA profiles were assessed by determining total (mg/ind) or relative (%) FA concentrations.

One‐way analysis of similarity (ANOSIM) was applied to determine differences in FA profiles of each species across the different treatments.

#### *2.2.5. Fatty acid trophic markers*

FA ratios of bacteria, algae or animal were assessed at the extracts of lipids of *C. edule*. The FA ratios determined and respective food sources are described in **Table 1**.

Fatty Acids' Profiles of Aquatic Organisms: Revealing the Impacts of Environmental and Anthropogenic Stressors http://dx.doi.org/10.5772/intechopen.68544 97


**Table 1.** Dietary and trophic fatty acid markers used in the present study.

### **3. Results**

solutions every next day. Bivalves were checked daily for mortality and behavioural conditions (to evaluate the conditions of the valves, organism behavior during feeding and the activity of the siphon). After the exposure period, all survival organisms were dissected, measured the weight and the body length and evaluated the condition indices of each individual. After the measurements, the muscle tissue (foot) was stored at −80°C for further fatty acid analysis.

The sample extraction for the FA analysis was obtained accordingly to the method described

PUFA detection [47]. A differentiated phase was extracted, and an internal standard (fatty acid Methylnonadecanoate (C19) Fluka 74208) was added to the quantification of FA. The samples were later analyzed using a gas chromatograph with a mass spectrometer and an HP88 column (60 m × 25 mm × 0.20 μm). It was conducted in splitless mode, with a 1 μL injector per run. The column temperature was set to increase from 75 to 230°C at a rate of 2°C/min. The carrier gas was helium at a flow rate of 1.3 mL/min. The results of the GC analyses were obtained, and fatty acid methyl esters (FAMEs) were identified by compari‐ son of their retention times with those of individual purified standards. FAMEs can also be quantified by determining the area of the peaks of each fatty acid with the help of calibration

The cellular density of *T. weissflogii* measured with a Neubauer Haemocytometer, at the end of the bioassays, was used to estimate the concentrations which induced *x*% growth inhibition

The LC10, LC20 and LC50 values with corresponding 95% confidence intervals for *C. edule*

To determine significant differences between treatments, one‐way analysis of variance (ANOVA) was performed, followed by Dunnett's multiple comparison test to identify signifi‐ cant differences between salinity treatments and the control treatment, considering a level of

The FA profiles were assessed by determining total (mg/ind) or relative (%) FA concentrations.

One‐way analysis of similarity (ANOSIM) was applied to determine differences in FA profiles

FA ratios of bacteria, algae or animal were assessed at the extracts of lipids of *C. edule*. The FA

ratios determined and respective food sources are described in **Table 1**.

regression, using the least‐squares method to fit the data to the logistic equation.

values, with *x* = 10, 20, 50) and the corresponding 95% confidence intervals by non‐linear

S0<sup>4</sup>

, due to reported deficiency in

in Gonçalves et al. [19], substituting BF3‐methanol by H<sup>2</sup>

*2.2.3. Fatty acid analysis*

96 Fatty Acids

factors [47].

(EC*<sup>x</sup>*

*2.2.4. Statistical analysis*

significance of 0.05.

*2.2.5. Fatty acid trophic markers*

were determined using Probit analysis [48].

of each species across the different treatments.

#### **3.1. Bioassays**

#### *3.1.1. Thalassiosira weissflogii*

After 96 h of exposure to a range of salinity concentrations and three different temperatures, significant statistical differences were observed between the control and the lower salinity concentrations with an exception to the bioassay conducted at 20°C where a significant sta‐ tistical difference was also observed to the highest salinity treatment (**Figure 2**). At the lowest temperature (15°C), the microalga presented the lowest growth than at higher temperatures. Still, to salinities near the optimal value of salinity to this microalga (30), the growth was high at all temperatures tested, not observing statistical significant differences between those con‐ centrations and the control (**Figure 2**).

Considering the exposure to both contaminants (the herbicide and the metal), *T. weissflogii* showed to be more sensitive to Primextra®Gold TZ than to copper (**Figure 3**).

A significant growth inhibition was detected after the exposure to both contaminants with the herbicide revealing to be more toxic than the metal. In fact, all treatments of the herbicide showed statistical significant differences with the control, whereas to copper, only the three higher concentrations presented statistical significant differences with the control.

#### *3.1.2. Cerastoderma edule*

Considering the optimality of salinity for the activity of *C. edule* is 20–25, the species revealed to be mostly affected by low salinities (LC50 = 11.01 (10.66–11.54) mg/L) with 100% of mortality at salinity concentrations below 10. Although the growth inhibition of *T. weissflogii* to lower salinity concentrations, the microalga demonstrated to be more tolerant than *C. edule*.

**Figure 2.** Cell density of *T. weissflogii* at (a) 15, (b) 20 and (c) 25°C after 96 h of exposure to salinity treatments, where CTL refers to the negative control treatment. Symbol '\*' indicates a significant (P < 0.05) difference of the treatments compared to the CTL.

**Figure 3.** Selected EC*<sup>x</sup>* (*x* = 10, 20, 50) values (mg/L‐1) estimated for *Thalassiosira weissflogii*. The central band of each box denotes the ECX (x=10, 20, 50) value and the bottom and top of the box represent the lower and upper 95% confidence limits.

In **Table 2**, there are the lethal concentration (LC) values determined to *C. edule* exposed to the herbicide and the metal. The results clearly revealed that estuarine bivalve is more sensitive to the metal than to the herbicide, showing an opposite pattern of the one observed with *T. weissflogii*.


**Table 2.** Lethal concentration (LC) values to *C. edule* exposed to copper sulfate and Primextra® Gold TZ. In brackets are indicated the 95% confidence limits.

#### **3.2. Fatty acid profiles**

#### *3.2.1. Thalassiosira weissflogii*

#### *3.2.1.1. Salinity experiments*

In **Table 2**, there are the lethal concentration (LC) values determined to *C. edule* exposed to the herbicide and the metal. The results clearly revealed that estuarine bivalve is more sensitive to the metal than to the herbicide, showing an opposite pattern of the one observed with *T. weissflogii*.

denotes the ECX (x=10, 20, 50) value and the bottom and top of the box represent the lower and upper 95% confidence

EC10 EC20 EC50 EC10 EC20 EC50

0.000 0.002 0.004 0.006 0.008 0.010 0.012

(*x* = 10, 20, 50) values (mg/L‐1) estimated for *Thalassiosira weissflogii*. The central band of each box

Cellular Density (cells/mL)

0 1e+5 2e+5 3e+5 4e+5 5e+5 6e+5 7e+5

Salinity Concentrations CTL 0 5 10 15 20 25 33

**Figure 2.** Cell density of *T. weissflogii* at (a) 15, (b) 20 and (c) 25°C after 96 h of exposure to salinity treatments, where CTL refers to the negative control treatment. Symbol '\*' indicates a significant (P < 0.05) difference of the treatments

\*

\* \*

**B**

Salinity Concentrations CTL 0 5 10 15 20 25 33

Primextra Gold TZ

\* \*

\*

\*

\*

Salinity concentrations CTL 0 5 10 15 20 25 33

Cellular Density (cells/mL)

0

Copper sulphate

2e+5

\*

\*

4e+5

6e+5

8e+5

**C**

\*

\*

\*

Cellular Density (cells/mL)

0.0 5.0e+4 1.0e+5 1.5e+5 2.0e+5 2.5e+5 3.0e+5 3.5e+5

compared to the CTL.

ECx (x=10, 20, 50) (mg/L)

0.0 0.1 0.2 0.3 0.4 0.5 0.6

**Figure 3.** Selected EC*<sup>x</sup>*

limits.

98 Fatty Acids

**A**

The fatty acid composition (mg/individual) of *T. weissflogii* exposed to three salinity treatments (10.46, 15.9, 18.5) plus the control is summarized in **Table 3** (15°C), **Table 4** (20°C) and **Table 5** (25°C). After the saline exposure to 15°C, the FA composition of *T. weissflogii* is varied in general with a significant decrease in MUFA and PUFA, especially at the salinity treatments of 10.46 and 15.9. The amount of SFAs in the control treatment is very low compared to other treatments. The saline treatment of 18.5 registered a drastic increase in the abundance of SFA compared to all treatments, with the control registering the lowest amount of saturated FA. MUFA was the domi‐ nant FA group in the FA profile of the marine diatom, with a decrease at the saline treatment of C2 (15.9), followed by C1 (18.5) and then C3 (10.46). In fact, the control at 15°C was the treatment that registered the highest amount in MUFA from the three temperatures tested. A dominance of LC‐MUFAs is notorious, although C20:1n9(cis11) was absent at the control and at the concen‐ tration C1 (18.5). PUFA also displayed a high abundance at the control and at C1 (18.5), with a decrease at the concentrations C2 (15.9) and C3 (10.46). In fact the total amount of PUFA at C1 treatment was almost twofold of the quantity registered at the control. Important precursors of LC‐PUFA such as C18:2n6c were presented only in the control and in the salinity concentration of 15.9, whereas C18:3n3 was presented in all treatments, with a residual quantity (closely zero) at the salinity treatment of 10.46. This FA group showed to be the most sensitive to salinity. HUFAs were presented at highest quantity at the extreme salinity treatments (C1 = 18.5; C3 = 10.46) with the control and the C2 treatment (15.9) registering the lowest values. All EFAs (ARA, EPA, DHA) had a similar pattern after saline exposure: their level remained very close in each concentration apart from EPA that registered the lowest value of the HUFA determined at all treatments. DHA was the most abundant EFA in the control treatment and at the salinity treatment of 18.5.

At 20°C, the FA profile of *T. weissflogii* exposed to the three saline treatments (15, 30.5 and 32) plus the control reported an increase in SFA content from the control to the lowest salin‐ ity concentration (15), although with relatively low diversity, had been notorious for higher amount of SFA of longer chain (**Table 4**). A significant rise was observed at the PUFA and HUFA contents at the lowest salinity treatment (15). An opposite trend was identified at the total amount of MUFA to all tested treatments when compared to the control. The salinity



**CTL**

> C14:0

C15:0

C16:0 C17:0

C18:0

C20:0 C21:0 C22:0

C23:0 C24:0 **Total SFA**

C14:1 C15:1

C16:1 C17:1 C18:1n9t C18:1n9c C20:1n9(cis‐11)

C22:1n9 C24:1n9 **Total MUFA**

C18:2n6t C18:2n6c

21.1455

 5.12853

212.8497

 26.0495

 400.9721

 79.7362

 19.2169 73.1763

 30.7402

 8.0727

35361.1083

 1989.1999

 18904.0593

 4093.5942

 19944.8750

 8231.0337

 1501.1175

6.9028

2.9890

 434.7142

316.5629

 39.3603

 425.4274

 169.0477

 139.9360

 58.7849

 116.9618

 29.7734

82.6656

 20.0494

 570.6772

 172.9728

 84.4090

 10.3481

 484.0046

 109.6170

48.2467

 0.3976

34.7431

 1.4553

49.0293

 2.9871

34777.3500

 1912.2145

 15868.4131

1262.8013

359.6901

417.0501

 117.3960

 45.6232 28.5713

 12.0023

 255.5463

 110.6548

 4.9456

 93.3756

 36.7104

 8.8740

 339.8028

 10046.7004

 4146.2759

 427.3974

2.1760 215.0315

 84.0745

0.9422

 99.6523

 3200.9993

 9562.9247

 3989.8029

52.5108

 12.7357

108.3361

 26.2754

 3269.1270

 1142.4632

 500.14525

 101.5802

 220.8601

 48.9027

75.8923

 18.4066

 395.4571

 157.1387

808.2884

803.2505

1207.2149

 479.6986

 162.62326

 467.5087

 87.8701

 195.8909

 38.09074

 321.1812

24.9692

 10.8120

32.4438

 7.8688

32.6365

 13.7101

**±Std. error**

 **C1 (18.5)**

54.9161

 21.8215

 **±Std. error**

 **C2 (15.9)**

 **±Std. error**

 **C3 (10.46)**

 **±Std. error**

100 Fatty Acids

**Table 3.** Abundance of fatty acids (saturated fatty acids—SFA, monounsaturated fatty acids—MUFA, polyunsaturated fatty acids—PUFA and highly unsaturated fatty acids—HUFA, in mg/ind) in the profile of *T. weissflogii* after exposure to salinity treatments (10.46; 15.9; 18.5) at 15°C.



**CTL**

> C14:0

C15:0

C16:0 C17:0

C18:0

C20:0 C21:0 C22:0 C23:0 C24:0 **Total SFA**

C14:1

C15:1

C16:1 C17:1 C18:1n9t C18:1n9c C20:1n9(cis‐11)

C22:1n9 C24:1n9 **Total MUFA**

C18:2n6t C18:2n6c

201.34907

 58.6126

 67.9254

 58.61256

 32.2866

 1.7356

32707.2860

 621.5169

 10945.4313

 621.5169

 28461.1817

 1964.4245

 3459.9672

198.9214

 59.3448

 1227.1800

14.1459

 7.0729

176.2199

 7.9133

386.1857

 112.0796

 76.3323 95.0662 278.4564

238.0316

 7.0729

 7.9133

78.0943

 2.8059

 112.0796

 25.4705

 1.9659

89.0514

 13.4534

 27.1444

 13.4534

 8.4810 58.9215

 13.1667004

 197.3032 390.8133

1704.3170

989.9304

156.5543

 59.1720

 324.5449

 629.4505

 131.4833

 74.5736

32041.6830

 480.9976

 10230.4003

 480.9976

 28290.2142

 1943.4874

2.9985

21.0489

 7.9558

143.3628

 48.8636

 164.2909

 48.8636

 206.3721

 51.5052

 2240.7950

 826.6445

67.8728

 21.6031

 45.6180 10.0689

12.5133

 4.4241

 21.6031

31.5296

 15.7648

 45.9347

 15.7648

 8.3243

2.9431

6.1552 35.3845

60.0374

 22.6920

 13.3741

2.3265

43.9605

 11.4958

 51.9012

 11.4958

10.7682

**±Std. error**

 **C1 (32)**

 **±Std. error**

 **C2 (30.5)** 185.5345

 44.1380

 1299.3503

171.8261

143.5637

 33.9657

 64.9442

 491.1083

 **±Sth. error**

 **C3 (15)** 524.4776

 198.2339

 **±Std. error**

102 Fatty Acids

**Table 4.** Abundance of fatty acids (saturated fatty acids—SFA, monounsaturated fatty acids—MUFA, polyunsaturated fatty acids—PUFA and highly unsaturated fatty acids—HUFA, in mg/ind) in the profile of *T. weissflogii* after exposure to salinity treatments (15; 30.5; 32) at 20°C.



**CTL**

> C14:0

C15:0 C16:0 C17:0

C18:0 C20:0 C21:0 C22:0 C23:0 C24:0 **Total SFA**

C14:1

C15:1

C16:1 C17:1

C18:1n9t C18:1n9c C20:1n9(cis‐11)

C22:1n9 C24:1n9 **Total MUFA**

C18:2n6t C18:2n6c

89.4747

 26.9962

 261.2764

 33.1211

4363.7445

 1877.394

 1880.3334

 518.1971

 3415.7770

 5184.6099

 77.1268

 14.7068

136.0564

 41.155

283.4383

 26.872

 23.9182

 4.8087

58.3992

106.4004

56.05451

1218.6964

440.7829

 105.7048

 295.9116

 31.2481

 9.9562

77.1268

 11.6267

 17.2988

 15.3800

 37.1824

 22.76945

 18.0224

3920.3316

 1557.7818

10178.9007

 3313.9257

666.8180

 132.0393

 2619.2930

 743.78106

 160.0438

 93.1342

 15.8881

 11.23460

324.9469

144.2169

 32.1972

 59.3290

 716.0168

 156.4173

 16.1034

 4.9800

133.0046

 21.8387

 301.2867

 92.9791

34.1393

 9.4712

14.7009

 2.9211

15.8093

 6.2820 265.4131

56.2356

51.8891

58.1041

 17.9313

 52.5980

 32.2000

 15.8881

 11.2346

 16.0133

 17.3547

 81.9083

 91.3424

 55.9356

**±Std. error**

 **C1 (23.76)** 1170.3476

 361.1771

 **±Std. error**

 **C2 (22.59)**

 **±Std. error**

 **C3 (14.79)**

 **±Std. error**

104 Fatty Acids

**Table 5.** Abundance of fatty acids (saturated fatty acids—SFA, monounsaturated fatty acids—MUFA, polyunsaturated fatty acids—PUFA and highly unsaturated fatty acids—HUFA, in mg/ind) in the profile of *T. weissflogii* after exposure to salinity treatments (14.79; 22.59; 23.76) at 25°C. concentration C2 (30.5) registered a sharp decrease in the total amount of MUFA, PUFA and HUFA when compared to the control. MUFA constituted the most representative group with the highest amount of FA in each treatment.

Comparing to the salinity experiments conducted at 15°C, the control treatment at 20°C pre‐ sented the highest content of SFA and PUFA, with the control at 15°C registered a higher amount of total MUFA and HUFA. Comparing the salinity treatments tested, the highest quantity of SFA, PUFA and HUFA was observed at the salinity concentrations C1 (18.5) and C3 (15) to the experiments performed at the temperature of 15 and 20°C, respectively. The total MUFA registered the highest values at the middle concentrations (C2 = 15.9; C2 = 30.5) of the experiments conducted at 15 and 20°C correspondingly.

At the FA profiles of *T. weissflogii* exposed to the salinity treatments at 25°C, a sharp decreased of total SFA from the control to the other treatments was observed with the exception of salinity concentration of 23.76, where a great increase was observed (**Table 5**). In general, MUFA constituted the most abundant group of FA, mainly represented by longer chain MUFAs, except at the highest salinity treatment (23.76). Still this salinity concentration registered the highest quantity of PUFA from all salinity experiments conducted at dis‐ tinct temperatures (15, 20 and 25°C). PUFA content was only observed in the control and at salinity concentration C3 (23.76). C18:3n3 was present in the higher salinity treatment and in low amounts at the control, which may indicate that these LC‐PUFA precursors were desaturated and elongated in the synthesis of HUFA. PUFA was characterized by a great diversity, still absent at the lower salinity treatments (14.79 and 22.59). HUFA registered the highest amount in the highest saline concentration (23.76) followed by the control treat‐ ment, decreased drastically at the other salinity treatments. Also, the control that registered the highest amount of HUFA was the one performed at 25°C. EFAs are mainly represented by ARA that were identified at all treatments. In the negative control, ARA is the most abun‐ dant EFA followed by EPA and lastly by DHA. Indeed, EPA and DHA were absent in the two lowest salinity treatments.

The FA content of *T. weissflogii* at the different salinity concentrations plus the control showed sharp changes among all treatments at 25°C than at the other temperatures (15 and 20°C). Furthermore, a great decrease in FA diversity was observed in C2 and C3 salinity treatments at the experiment conducted at 25°C, whereas the experiments conducted at 15 and 20°C pre‐ sented higher or similar diversity of FA among the treatments, which indicate a detrimental effect of salinity in the FA content of the species.

#### *3.2.1.2. Pollutants experiments*

The results obtained demonstrated the fatty acid profiles of the microalga were affected by the presence of both toxicants, mainly by the metal (**Table 6**). Although it was not detected clear differences among the treatments, moderate changes were observed at the highest herbicide concentration compared to the other treatments. Small changes at the total amount of SFA and MUFA were also registered between the control and the highest concentration (0.0078 mg/L) of Primextra.

Fatty Acids' Profiles of Aquatic Organisms: Revealing the Impacts of Environmental and Anthropogenic Stressors http://dx.doi.org/10.5772/intechopen.68544 107


**Table 6.** Total fatty acid and EFA (%) content in *T. weissflogii* after the exposure to the herbicide Primextra® Gold TZ and the metal copper.

A marginal decrease of SFA from the CTL to the copper concentration of 0.1995 mg/L, with a rise at the highest copper treatment was observed. An opposite trend was verified to the unsaturated fatty acids (MUFA, PUFA and HUFA). A slightly rise in the total amount of the UFA was registered from the CTL to the metal concentration of 0.1995 mg/L followed by a decrease at the highest treatment. Although significant differences were not registered at the fatty acid content between the control and the lowest copper treatments, clear differences between the control and the highest copper concentrations were verified.

The amount of HUFA in herbicide and copper treatments was related to the presence of DHA and EPA, with ARA being absent at all treatments. EPA was the dominant EFA at all treatments (**Table 6**).

#### *3.2.2. Cerastoderma edule*

concentration C2 (30.5) registered a sharp decrease in the total amount of MUFA, PUFA and HUFA when compared to the control. MUFA constituted the most representative group with

Comparing to the salinity experiments conducted at 15°C, the control treatment at 20°C pre‐ sented the highest content of SFA and PUFA, with the control at 15°C registered a higher amount of total MUFA and HUFA. Comparing the salinity treatments tested, the highest quantity of SFA, PUFA and HUFA was observed at the salinity concentrations C1 (18.5) and C3 (15) to the experiments performed at the temperature of 15 and 20°C, respectively. The total MUFA registered the highest values at the middle concentrations (C2 = 15.9; C2 = 30.5) of

At the FA profiles of *T. weissflogii* exposed to the salinity treatments at 25°C, a sharp decreased of total SFA from the control to the other treatments was observed with the exception of salinity concentration of 23.76, where a great increase was observed (**Table 5**). In general, MUFA constituted the most abundant group of FA, mainly represented by longer chain MUFAs, except at the highest salinity treatment (23.76). Still this salinity concentration registered the highest quantity of PUFA from all salinity experiments conducted at dis‐ tinct temperatures (15, 20 and 25°C). PUFA content was only observed in the control and at salinity concentration C3 (23.76). C18:3n3 was present in the higher salinity treatment and in low amounts at the control, which may indicate that these LC‐PUFA precursors were desaturated and elongated in the synthesis of HUFA. PUFA was characterized by a great diversity, still absent at the lower salinity treatments (14.79 and 22.59). HUFA registered the highest amount in the highest saline concentration (23.76) followed by the control treat‐ ment, decreased drastically at the other salinity treatments. Also, the control that registered the highest amount of HUFA was the one performed at 25°C. EFAs are mainly represented by ARA that were identified at all treatments. In the negative control, ARA is the most abun‐ dant EFA followed by EPA and lastly by DHA. Indeed, EPA and DHA were absent in the

The FA content of *T. weissflogii* at the different salinity concentrations plus the control showed sharp changes among all treatments at 25°C than at the other temperatures (15 and 20°C). Furthermore, a great decrease in FA diversity was observed in C2 and C3 salinity treatments at the experiment conducted at 25°C, whereas the experiments conducted at 15 and 20°C pre‐ sented higher or similar diversity of FA among the treatments, which indicate a detrimental

The results obtained demonstrated the fatty acid profiles of the microalga were affected by the presence of both toxicants, mainly by the metal (**Table 6**). Although it was not detected clear differences among the treatments, moderate changes were observed at the highest herbicide concentration compared to the other treatments. Small changes at the total amount of SFA and MUFA were also registered between the control and the highest concentration (0.0078 mg/L)

the highest amount of FA in each treatment.

106 Fatty Acids

two lowest salinity treatments.

*3.2.1.2. Pollutants experiments*

of Primextra.

effect of salinity in the FA content of the species.

the experiments conducted at 15 and 20°C correspondingly.

#### *3.2.2.1. Salinity experiments*

A considerable increase in the main FA groups (saturated fatty acids—SFA and unsaturated fatty acids—UFA) was clearly observed in the individuals from the field to the organisms exposed to a range of salinity concentrations, unless to polyunsaturated fatty acid (PUFA) where an opposite trend was verified (**Table 7**). Individuals from the field were mostly con‐ stituted by PUFA (95.773%), presented lower amounts of SFA (3.314%), HUFA (0.708%) and MUFA (0.205%). Under a range of salinity concentrations, a lacking of SFA of short chain was clearly observed in all organisms, with only the individuals exposed to the highest salinity treatment presented slightly amounts of C6:0, C8:0 and C10:0. Omega‐6 was mainly repre‐ sented by γ‐linolenic acid, C18:3n6 and arachidonic acid (ARA), C20:4n6, whereas omega‐3 occurred mainly at the forms of docosahexaenoic acid (DHA), C22:6n3, eicosapentaenoic acid (EPA), C20:5n3 and α‐Linolenic acid (ALA), C18:3n3.


**Table 7.** Total fatty acid and EFA (%) content in *C. edule* in the field and after the exposure to a range of salinity treatments.

#### *3.2.2.2. Pollutants experiments*

A slightly increase at all FA groups was observed in the organisms exposed to the herbicide com‐ pared with the individuals from the field (**Table 8**). An opposite pattern was found in the organ‐ isms exposed to copper where a slender decrease in SFA (at the first metal treatment), PUFA and HUFA was observed compared to the individuals from the field. Comparing the HUFA content in the organisms exposed to both pollutants, a slightly increase was observed at the herbicide treatments with a reduction at the copper concentrations. DHA was the EFA occurring in higher amount after the exposure to commercial formulation of the herbicide. In the treatments exposed to the metal, a clear pattern was not observed. In general, there was a higher diversity in FA in the individuals exposed to Primextra compared to the organisms under copper treatments, with exception to the highest copper concentration that registered the highest diversity of FA (**Table 8**).


**Table 8.** Total fatty acid and EFA (%) content in *C. edule* in the field and after the exposure to a set concentrations of the herbicide Primextra® Gold TZ and the metal copper.

#### *3.2.2.3. Fatty acid trophic markers (FATMs): Cerastoderma edule*

In **Table 9**, the FA composition of the food (rotifers and microalgae) used daily to feed *C. edule* in the lab is represented. Microalgae food source presented a highest richness in PUFA whereas rotifers showed higher composition in SFA.

FATM ratios indicated an omnivorous diet with the organisms ingesting phytoplankton and zooplankton, with some of the individuals in the lab and at the field consuming higher amounts of phytoplankton (diatoms). Still some organisms from the field also showed a diet based mainly on zooplankton with few also feeding on bacteria.


**Table 9.** Fatty acid composition of food source used daily to feed *Cerastoderma edule* in the lab.

### **4. Conclusion**

*3.2.2.2. Pollutants experiments*

% total SFA

108 Fatty Acids

% total MUFA

% total PUFA

% total HUFA

herbicide Primextra® Gold TZ and the metal copper.

**Salinity concentrations**

A slightly increase at all FA groups was observed in the organisms exposed to the herbicide com‐ pared with the individuals from the field (**Table 8**). An opposite pattern was found in the organ‐ isms exposed to copper where a slender decrease in SFA (at the first metal treatment), PUFA and HUFA was observed compared to the individuals from the field. Comparing the HUFA content in the organisms exposed to both pollutants, a slightly increase was observed at the herbicide treatments with a reduction at the copper concentrations. DHA was the EFA occurring in higher amount after the exposure to commercial formulation of the herbicide. In the treatments exposed to the metal, a clear pattern was not observed. In general, there was a higher diversity in FA in the individuals exposed to Primextra compared to the organisms under copper treatments, with exception to the highest copper concentration that registered the highest diversity of FA (**Table 8**).

**Primextra® Gold TZ (mg/L) Copper (mg/L)**

**Field CTL 0.5 2.5 5 10 20 30 Field CTL 0.6 0.9 1.2**

0.074 0.129 0.204 0.0744 0.155 0.130 0.165 0.098 0.016 0.001 0.005 0.052 0.077

0.019 0.041 0.057 0.023 0.039 0.036 0.039 0.023 0.025 0.030 0.029 0.029 0.031

0.051 0.108 0.061 0.049 0.066 0.054 0.069 0.048 0.153 0.042 0.078 0.086 0.121

0.113 0.187 0.258 0.110 0.221 0.196 0.224 0.150 0.116 0.084 0.159 0.144 0.075

*EPA* 0.047 0.084 0.095 0.050 0.069 0.075 0.070 0.062 0.084 0.042 0.094 0.074 0.034 *DHA* 0.067 0.102 0.164 0.061 0.156 0.121 0.154 0.088 0.032 0.041 0.065 0.070 0.040 *N* 11 12 15 12 11 13 13 12 11 9 12 12 16

**Table 8.** Total fatty acid and EFA (%) content in *C. edule* in the field and after the exposure to a set concentrations of the

**Table 7.** Total fatty acid and EFA (%) content in *C. edule* in the field and after the exposure to a range of salinity treatments.

**Field 10 15 25 30 35**

% total SFA 3.314 26.561 31.434 19.843 59.711 30.511 % total MUFA 0.205 32.280 28.336 15.043 14.606 19.847 % total PUFA 95.773 25.130 16.162 44.712 4.095 42.821 % total HUFA 0.708 16.030 24.068 20.401 21.588 6.821 *ARA* 0.000 0.000 3.849 5.401 9.963 3.245 *EPA* 0.117 14.835 15.468 7.888 7.838 2.988 *DHA* 0.591 1.195 4.750 7.112 3.786 0.588 *N* 28 21 15 15 15 22

> In general, an organism under stress conditions may change its physiological and biochemical responses as a strategic mechanism to compensate the organism's requirements [20]. In this study, the species from different trophic levels (a marine microalgae and estuarine bivalve spe‐ cies) exposed to a range of salinity concentrations under distinct temperature conditions and under different treatments of a metal (copper) and a herbicide (Primextra) revealed changes in its fatty acid content. The results obtained confirmed that the environmental and chemical stressors affect the fatty acid profile of aquatic species with sharp changes in the FA content of these species and reflecting then in lower quality food. It was clear that higher temperature had a great impact on the FA composition of the microalga, with the diatom not presenting several FA mainly PUFA, DHA and EPA in its profile, making the microalga more vulnerable to the effects of different salinities, mainly under salinity concentrations lower than 22.59. In the salinity treatments, an increase of SFA and MUFA was also observed to both studied spe‐ cies which can indicate that PUFA and HUFA are being metabolized so that the cells can obtain more energy necessary to maintain homeostatic ionic balance while maintaining basal func‐ tions of the body, such as respiration and excretion of ammonia. Since synthesizing of PUFAs and HUFAs becomes too energetically costly, the organism may not complete the elongation processes presenting higher concentrations of SFAs and MUFAs in the FA composition. The increase in saturation level can also be explained by a cellular response toward the osmotic shock, an attempt to maintain the stability of lipid membranes [49] and as such maintain the osmotic pressure from cell damaging. A low concentration of HUFA in general may also occur due to the high sensitiveness of this FA group to environmental fluctuations and to cell stress.

In literature studies, the most characteristic FA of diatoms are included in SFA and MUFA group and are characteristic of the plant domain: C14:0, C16:0, C16:1, C18:4 and C20:5, pre‐ cursors for LC‐PUFA such as EPA and DHA, while in turn most diatoms are reported to be very poor in C18:2 and C18:3 [50, 51]. In the present study, the absence of most of these FA at both species with a wide lacking of SFA is clearly shown. Fisher [52] stated *Thalassiosira pseudonana* Hasle & Heimdal was dominated by the FA reported above, although in differ‐ ent relative amounts. Fisher [52] also observed that the concentration of C16:0 varied with the culture age and that C16:1 declined drastically in the dark. The LC‐PUFA content of microalgae can indeed depend not only on the species, but also on factors related to cul‐ ture condition including composition of the medium, aeration, light intensity, temperature and age of culture [53], being crucial algae be maintained under its optimality of growth conditions.

In bivalve species, the reproductive success is related to the presence of great amount of lipids that are the second major constituent of bivalves' eggs [54, 55]. The maturation of the germ cell is closely related to the FA C18:3n3, C18:4n3, C20:1n9 and C20:2n6, while C14:0, C16:0, C16:1n7, C18:1n7 and C18:1n9 play a key role at the embryonic development of the bivalve eggs [54, 56]. Thus, it is crucial the presence of these FA during the breeding and thus in the FA profiles of the individuals.

The low concentration of HUFA reported in the lowest salinity treatments (22.59 and 14.79) at 25°C to *T. weissflogii* and to *C. edule* exposed to the metal may translate a metabolic response from the cells, which cannot efficiently maintain the cellular homeostatic ionic balance and cannot use energetic reservations in the elongation and synthesis of LC‐PUFA such as EPA (C20:5n3) and DHA (C22:6n3). In the salinity treatments, EFAs of both studied species (the diatom and the bivalve) were mainly represented by ARA that is an important precursor of signalling molecules including prostaglandins, prostacyclins and thromboxanes [57]. DHA was the EFA that presented the lowest concentration or was absent in all salinity treatments at the highest temperature (25°C) and also under the exposure to both pollutants of the diatom species. A similar pattern was reported to the bivalve species exposed to saline and copper concentrations. This EFA is crucial in neurophysiologic processes and influences visual acu‐ ity once is abundant in retina cells, as well as a preventive role in cardiac diseases being also presented in high amount in the brain tissues [4, 16]. In fact, a high concentration of EFA such as the combination DHA + EPA is linked to the reduction of coronary heart diseases. These two FA molecules also show a key role in bipolar disorder and other neurological dysfunc‐ tions, cognitive functions and fetal development. They are associated with benefits in the treatment of rheumatoid arthritis and inflammatory bowel disease, as well as for Crohn's disease, which is related with the suppression of ARA‐derived eicosanoids [4]. Once EPA and ARA play an important role in mediating immunological responses to infections and regulat‐ ing ion and water flux, a low or absent content of such EFAs can translate into alterations in membrane phospholipids once ARA and DHA components can influence cellular signalling but also sharply alter many membrane physical properties such as fluidity and bilayer thick‐ ness, among others. In humans, a deficiency of DHA affects neurotransmission, membrane‐ bound enzyme and ion channel activities, intensity of inflammation and immunity, all of these associated with normal aging, Alzheimer disease, hyperactivity and schizophrenia [13]. These EFAs (EPA and ARA) are also associated to the further improvement of adaption of the individuals to anthropogenic and environmental stress conditions [58]. Furthermore, C18:3n3 (α LNA) is associated with the processes of elongation and desaturation at the synthesis of EPA and DHA in mammals, whereas C18:2n6, C20:4n6 and C18:3n6 are involved in the bio‐ synthesis of PUFA of long chain [59].

The level of essential fatty acids (n‐3 EFA) in algae can be highly variable, [60], including among major taxa [61], making it hard to compare between phylogenetic close species. PUFAs and HUFAs can reach their highest content during periods of rapid cell growth or bloom epi‐ sodes [62] and are important components of the microalgae membranes that affect cell mem‐ brane fluidity [3] that promotes a rapid response to environmental changes, such as variations in temperature, light and pH. Thus, HUFAs are the most affected FA group in case of cellular damage due to failed osmoregulation. The reduction of EFA content, mainly in the base of the trophic food web, may have serious implications at higher levels in the food web, once the compromising of the nutritious value of the primary producers influences the uptake of EFAs and thus the fundamental processes of regulation of many species in the ecosystems. Some of these species are the food source of human beings, having the climate events in larger scale a great impact at the biochemical values of nutritional requirements of many aquatic species and thus at human health. Thus, a balanced fatty acid profile is essential, with a balanced amount of essential fatty acids (EFAs) and of other fatty acids with a key role in the regulation and functioning of the organisms.

Fatty acid proves to be a useful bio‐indicator of ecological and healthy status of aquatic eco‐ systems, providing crucial information about the impacts of global stressors in the aquatic communities and thus in the trophic food web with severe repercussions to human beings and food quality. Recent reports predict the occurrence, at the next 100 years, of changes in salinity seawater, rise in temperature and water acidification [63]. In addition to these climate changes, an intensive agriculture production with an excessive usage of fertilizers and pesticides near coastal wetlands will have severe impacts to the aquatic communities and thus to the ecosystem. Therefore, it is of major importance and becomes a priority to determine and predict the effects of environmental and anthropogenic stressors in the aquatic systems in order to maintain the healthy status and the biodiversity and thus the food quality.

### **Acknowledgements**

In literature studies, the most characteristic FA of diatoms are included in SFA and MUFA group and are characteristic of the plant domain: C14:0, C16:0, C16:1, C18:4 and C20:5, pre‐ cursors for LC‐PUFA such as EPA and DHA, while in turn most diatoms are reported to be very poor in C18:2 and C18:3 [50, 51]. In the present study, the absence of most of these FA at both species with a wide lacking of SFA is clearly shown. Fisher [52] stated *Thalassiosira pseudonana* Hasle & Heimdal was dominated by the FA reported above, although in differ‐ ent relative amounts. Fisher [52] also observed that the concentration of C16:0 varied with the culture age and that C16:1 declined drastically in the dark. The LC‐PUFA content of microalgae can indeed depend not only on the species, but also on factors related to cul‐ ture condition including composition of the medium, aeration, light intensity, temperature and age of culture [53], being crucial algae be maintained under its optimality of growth

In bivalve species, the reproductive success is related to the presence of great amount of lipids that are the second major constituent of bivalves' eggs [54, 55]. The maturation of the germ cell is closely related to the FA C18:3n3, C18:4n3, C20:1n9 and C20:2n6, while C14:0, C16:0, C16:1n7, C18:1n7 and C18:1n9 play a key role at the embryonic development of the bivalve eggs [54, 56]. Thus, it is crucial the presence of these FA during the breeding and thus in the

The low concentration of HUFA reported in the lowest salinity treatments (22.59 and 14.79) at 25°C to *T. weissflogii* and to *C. edule* exposed to the metal may translate a metabolic response from the cells, which cannot efficiently maintain the cellular homeostatic ionic balance and cannot use energetic reservations in the elongation and synthesis of LC‐PUFA such as EPA (C20:5n3) and DHA (C22:6n3). In the salinity treatments, EFAs of both studied species (the diatom and the bivalve) were mainly represented by ARA that is an important precursor of signalling molecules including prostaglandins, prostacyclins and thromboxanes [57]. DHA was the EFA that presented the lowest concentration or was absent in all salinity treatments at the highest temperature (25°C) and also under the exposure to both pollutants of the diatom species. A similar pattern was reported to the bivalve species exposed to saline and copper concentrations. This EFA is crucial in neurophysiologic processes and influences visual acu‐ ity once is abundant in retina cells, as well as a preventive role in cardiac diseases being also presented in high amount in the brain tissues [4, 16]. In fact, a high concentration of EFA such as the combination DHA + EPA is linked to the reduction of coronary heart diseases. These two FA molecules also show a key role in bipolar disorder and other neurological dysfunc‐ tions, cognitive functions and fetal development. They are associated with benefits in the treatment of rheumatoid arthritis and inflammatory bowel disease, as well as for Crohn's disease, which is related with the suppression of ARA‐derived eicosanoids [4]. Once EPA and ARA play an important role in mediating immunological responses to infections and regulat‐ ing ion and water flux, a low or absent content of such EFAs can translate into alterations in membrane phospholipids once ARA and DHA components can influence cellular signalling but also sharply alter many membrane physical properties such as fluidity and bilayer thick‐ ness, among others. In humans, a deficiency of DHA affects neurotransmission, membrane‐ bound enzyme and ion channel activities, intensity of inflammation and immunity, all of

conditions.

110 Fatty Acids

FA profiles of the individuals.

This study was supported by the Fundação para a Ciência e Tecnologia (FCT) through the strategic projects UID/MAR/04292/2013 granted to MARE and UID/AMB/50017/2013 granted to CESAM. A.M.M. Gonçalves also thanks the Fundação para a Ciência e Tecnologia (FCT) for the financial support provided through the post‐doctoral grant SFRH/BPD/97210/2013 co‐funded by the Human Potential Operational Programme (National Strategic Reference Framework 2007–2013), European Social Fund (EU) and the program POPH/FSE.

### **Author details**

Ana M.M. Gonçalves1,2\*, João C. Marques<sup>1</sup> and Fernando Gonçalves<sup>2</sup>

\*Address all correspondence to: anamartagoncalves@ua.pt; anamartagoncalves@gmail.com

1 IMAR (Marine and Environmental Research Centre) & MARE (Marine and Environmental Sciences Centre), Faculty of Sciences and Technology, University of Coimbra, Coimbra, Portugal

2 Department of Biology and CESAM, University of Aveiro, Aveiro, Portugal

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**Author details**

Portugal

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## **Fatty Acids from Microalgae: Targeting the Accumulation of Triacylglycerides**

Paola Scodelaro Bilbao, Gabriela A. Salvador and Patricia I. Leonardi

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/67482

#### **Abstract**

Microalgae were originally considered as sources of long-chain polyunsaturated fatty acids (PUFAs), mainly for aquaculture purposes. However, based on the fact that their fatty acids (FA), stored as triacylglycerides (TAG), can be converted into biodiesel via a transesterification reaction, several microalgal species have emerged over the last decade as promising feedstocks for biofuel production. Elucidation of microalgae FA and TAG metabolic pathways is therefore becoming a cutting-edge field for developing transgenic algal strains with improved lipid accumulation ability. Furthermore, many of the biomolecules produced by microalgae can also be exploited. In this chapter, we describe recent advances in the field of FA and TAG pathways in microalgae, focusing in particular on the enzymes involved in FA and TAG synthesis, their accumulation in lipid droplets, and their degradation. Mention is made of potentially high-value products that can be obtained from microalgae, and possible molecular targets for enhancing FA and TAG production are outlined. A summary is provided of transcriptomics, proteomics, and metabolomics of the above-mentioned pathways in microalgae. Understanding the relation between anabolic and catabolic lipid enzyme pathways will provide new insights into biodiesel production and other valuable biomolecules obtained from microalgae.

**Keywords:** fatty acids, triacylglycerides, lipid metabolism, microalgae

### **1. Introduction**

Despite the drop in crude oil prices over the last few years, global efforts to develop alternative renewable energy sources continue to be driven by increasing air pollution and growing energy consumption. Extensive research is therefore being conducted in the field of biofuels

© 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

[1], which are derived from renewable biological sources. Biodiesel is the main substitute for diesel fuel and can be produced from both edible and non-edible oils. The use of edible oils has generated controversy because of the negative impact on food availability and the environment [2, 3]. As a consequence of these ethical considerations, non-food crops have emerged as a viable alternative for the production of biodiesel [4–6]. However, since non-food crops do not produce sufficient biomatter to feasibly cover the fuel requirements of the world's transport sector, attention is turning to oleaginous microalgae which are able to produce and accumulate large amounts of fatty acids (FA) in the form of triacylglycerides (TAG) that can be converted into biodiesel through a transesterification reaction [2, 3, 7]. Furthermore, some species of oleaginous microalgae can also produce high-value products such as long-chain polyunsaturated fatty acids (docosahexaenoic (DHA) and eicosapentaenoic (EPA) acids), carbohydrates (cellulose, starch), proteins, and other high-value compounds, such as pigments, antioxidants (i.e., β-carotene, astaxanthin), and vitamins, which may have commercial application in various industrial sectors [2, 3, 8, 9]. In addition to their potential as biological factories, the advantage of these photosynthetic microorganisms is that their simple growing requirements (light, CO2, and nutrients) offer several environmental benefits such as high solar energy conversion efficiency, utilization of saline water, CO<sup>2</sup> sequestration from the air and self-purification if coupled with wastewater treatment [10].

Despite the wide range of metabolites able to be synthesized by microalgae, little is known about the regulation of FA and TAG biosynthetic pathways and their storage and turnover in microalgae. In this chapter, we therefore describe recent advances in these fields and possible high-value co-products that could render the production of biodiesel from microalgae more sustainably. Recent studies on the transcriptomics, proteomics, and metabolomics of the abovementioned pathways are also outlined. Understanding these metabolic pathways will accelerate the availability of biodiesel and other valuable biomolecules obtained from microalgae.

### **2. FA and TAG biosynthetic pathways in microalgae**

Fatty acids are organic acids containing a carboxylic functional group with an aliphatic chain that can be saturated (SFA), monounsaturated (MUFA), or polyunsaturated (PUFA). The number of carbon atoms can vary, generating short-chain, medium-chain, or long-chain FA.

In plants, the FA biosynthetic pathway occurs in the chloroplasts (**Figure 1**).

As shown in **Figure 1**, the first step in the pathway involves the acetyl-CoA carboxylase (ACCase) which catalyzes the formation of malonyl-CoA from acetyl-CoA and bicarbonate [11]. There is evidence suggesting the presence of genes encoding this enzyme (accA and accD) in *Chlorella pyrenoidosa*. In fact, the transcription of these genes showed to be up-regulated under lipid accumulating conditions [12]. Moreover, a marked increase in the level of acetyl-CoA together with a moderate augmentation of malonyl-CoA and CoA was detected in the green microalgae *Chlorella desiccata, Dunaliella tertiolecta*, and *Chlamydomonas reinhardtii* under stress conditions, denoting increased activity of ACCase in these strains [13].

[1], which are derived from renewable biological sources. Biodiesel is the main substitute for diesel fuel and can be produced from both edible and non-edible oils. The use of edible oils has generated controversy because of the negative impact on food availability and the environment [2, 3]. As a consequence of these ethical considerations, non-food crops have emerged as a viable alternative for the production of biodiesel [4–6]. However, since non-food crops do not produce sufficient biomatter to feasibly cover the fuel requirements of the world's transport sector, attention is turning to oleaginous microalgae which are able to produce and accumulate large amounts of fatty acids (FA) in the form of triacylglycerides (TAG) that can be converted into biodiesel through a transesterification reaction [2, 3, 7]. Furthermore, some species of oleaginous microalgae can also produce high-value products such as long-chain polyunsaturated fatty acids (docosahexaenoic (DHA) and eicosapentaenoic (EPA) acids), carbohydrates (cellulose, starch), proteins, and other high-value compounds, such as pigments, antioxidants (i.e., β-carotene, astaxanthin), and vitamins, which may have commercial application in various industrial sectors [2, 3, 8, 9]. In addition to their potential as biological factories, the advantage of these photosynthetic microorganisms is that their simple growing requirements (light, CO2, and nutrients) offer several environmental benefits such as high

Despite the wide range of metabolites able to be synthesized by microalgae, little is known about the regulation of FA and TAG biosynthetic pathways and their storage and turnover in microalgae. In this chapter, we therefore describe recent advances in these fields and possible high-value co-products that could render the production of biodiesel from microalgae more sustainably. Recent studies on the transcriptomics, proteomics, and metabolomics of the abovementioned pathways are also outlined. Understanding these metabolic pathways will accelerate the availability of biodiesel and other valuable biomolecules obtained from microalgae.

Fatty acids are organic acids containing a carboxylic functional group with an aliphatic chain that can be saturated (SFA), monounsaturated (MUFA), or polyunsaturated (PUFA). The number of carbon atoms can vary, generating short-chain, medium-chain, or long-chain FA.

As shown in **Figure 1**, the first step in the pathway involves the acetyl-CoA carboxylase (ACCase) which catalyzes the formation of malonyl-CoA from acetyl-CoA and bicarbonate [11]. There is evidence suggesting the presence of genes encoding this enzyme (accA and accD) in *Chlorella pyrenoidosa*. In fact, the transcription of these genes showed to be up-regulated under lipid accumulating conditions [12]. Moreover, a marked increase in the level of acetyl-CoA together with a moderate augmentation of malonyl-CoA and CoA was detected in the green microalgae *Chlorella desiccata, Dunaliella tertiolecta*, and *Chlamydomonas reinhardtii* under

sequestration from the air

solar energy conversion efficiency, utilization of saline water, CO<sup>2</sup>

**2. FA and TAG biosynthetic pathways in microalgae**

In plants, the FA biosynthetic pathway occurs in the chloroplasts (**Figure 1**).

stress conditions, denoting increased activity of ACCase in these strains [13].

and self-purification if coupled with wastewater treatment [10].

120 Fatty Acids

**Figure 1.** Simplified overview of the pathways involved in FA synthesis in plants. Enzyme abbreviations: ACCase, acetyl-CoA carboxylase; MCAT, malonyl-CoA:Acyl Carrier Protein (ACP) transacylase; KAS, ketoacyl-ACP synthases.

The next step in the FA synthesis is mediated by the malonyl-CoA:Acyl Carrier Protein (ACP) transacylase (MCAT) which transfers the malonyl group from malonyl-CoA to malonyl-ACP [11]. A putative MCAT was identified as a part of the FA biosynthetic pathway in *Nannochloropsis oceanica* [14]. In *Haematococcus pluvialis*, the genes encoding ACP were up-regulated under TAG accumulating conditions (high temperature, high salinity, and nitrogen deficiency) together with other genes involved in FA biosynthesis [15]. In addition, proteomic studies on *Neochloris oleoabundans* revealed an augmented expression of ACP, among other enzymes of the lipid synthesis, under nitrogen starvation [16].

Acyl-ACP is the carbon source or substrate for the elongation of FA. This reaction is catalyzed by enzymes known as ketoacyl-ACP synthases (KASIII, KASI, and KASII). After each condensation, a reduction, dehydration, and second reduction occur. These steps are catalyzed by enzymes known as the FAS complex: beta-ketoacyl-ACP reductase (KAR), hydroxyacyl-ACP dehydrase (HAD), and enoyl-ACP reductase (EAR), respectively [11]. Transcriptome analysis of the diatom *Chaetoceros* sp. GSL56 helped to identify putative enzymes of the FA synthesis pathway. In addition, replacement of ketoacyl-ACP synthase of *Synechococcus* 7002 with *Chaetoceros* ketoacyl-ACP synthase III induced FA synthesis [17]. In line with this, TAG accumulating conditions increased the levels of transcripts for KAS in *H. pluvialis* [15].

The *de novo* resulting FA often with 16 or 18 carbon atoms can undergo the action of elongases and desaturases that add carbon or double bonds, respectively [11]. Particularly, desaturases and elongases are being intensively studied to achieve transgenic long-chain PUFA production [18, 19].

Some reports suggest the presence of both enzyme types in microalgae. In the marine microalgae *Pavlova* sp. and *Isochrysis sp.*, two genes encoding elongases that catalyze the elongation of eicosapentaenoic acid (EPA) to docosahexaenoic acid (DHA) have been reported [20]. In the diatom *Thalassiosira pseudonana*, the genes encoding elongases that mediate the formation of DHA from EPA were successfully overexpressed, thus inducing an increase in DHA content [19]. A delta 5 desaturase was also identified, characterized and overexpressed in the diatom *Phaeodactylum tricornutum* inducing a significant increase in the unsaturated fatty acids [21] .

Upon completion of elongation, FAs are transported to the cytoplasm to act as substrates of the acyl transferases involved in the TAG synthesis. TAG are neutral lipids formed by the esterification of one molecule of glycerol with three FAs. Because of their energy-rich acyl chains, they are the dominant form of stored energy in microalgae. Cellular stresses, such as nutrient deprivation (carbon dioxide, nitrogen, silica, and phosphorous), temperature fluctuation, or high light exposure trigger their formation [22–28]. It has been demonstrated that lipid biosynthetic pathways are induced under these conditions to potentiate the lipid storage (30–60% of dry cell weight), and this mechanism is thought to play a role in microalgae adaptation and survival [24, 29–39]. It has further been reported that multiple stressors have no additive effect on lipid accumulation [24, 40].

Data relating to plant FA and TAG metabolism provided the key to identifying possible molecular targets involved in lipid synthesis and accumulation in microalgae [41]. As shown in **Figure 2**, in plants, the first step of the conventional Kennedy pathway involves the acylation of the glycerol-3-phosphate (G-3-P), catalyzed by the glycerol-3-phosphate acyltransferase (GPAT) to yield lysophosphatidic acid (LPA). GPAT is the rate-limiting step subject to many regulatory controls at the transcriptional and post-transcriptional level and to allosteric mechanisms [42, 43]. Recent studies have revealed the presence of this enzyme in microalgae. In the marine diatom *T. pseudonana*, a membrane-bound GPAT designated TpGPAT was cloned and characterized. The authors observed that G-3-P was the preferred substrate of TpGPAT [44]. A sequence for GPAT with high homology to that of plants was found in *C. reinhardtii, Volvox carteri, Ostreococcus lucimarinus, Ostreococcus tauri, Cyanidioschyzon merolae*, and *P. tricornutum*. As in *T. pseudonana*, G-3-P and fatty acyl molecules are likely to be the enzyme substrates, as suggested by the residues present in their active sites [45].

As described in **Figure 2**, lysophosphatidic acid acyltransferase (LPAAT) participates in the second step of the Kennedy pathway. This enzyme catalyzes the acylation of the LPA to yield phosphatidic acid (PA) [46]. Candidate LPAATs have been found in some algal genomes including that of *H. pluvialis* [47, 48], where it has been shown that LPAAT mRNA is induced under high irradiance stress [47]. In addition, it was recently reported that the expression of *C. reinhardtii* LPAAT (CrLPAAT1) is associated with an increase in lipid synthesis and accumulation under nitrogen starvation [48].

Phosphatidic acid phosphohydrolase (PAP) uses PA as substrate to form diacylglycerol (DAG), a precursor of TAG (**Figure 2**) [49]. In eukaryotes, PAP enzymes are the members of the evolutionarily conserved lipin protein family whose activity is related to TAG storage [50]. In the green microalga *C. reinhardtii*, PAP transcripts (named CrPAP2) are induced under stress conditions. In addition, CrPAP2 silencing slightly lowers the lipid content. Thus, in *C. reinhardtii*, as in other eukaryotes, PAP expression is related to lipid synthesis and accumulation [49].

Some reports suggest the presence of both enzyme types in microalgae. In the marine microalgae *Pavlova* sp. and *Isochrysis sp.*, two genes encoding elongases that catalyze the elongation of eicosapentaenoic acid (EPA) to docosahexaenoic acid (DHA) have been reported [20]. In the diatom *Thalassiosira pseudonana*, the genes encoding elongases that mediate the formation of DHA from EPA were successfully overexpressed, thus inducing an increase in DHA content [19]. A delta 5 desaturase was also identified, characterized and overexpressed in the diatom *Phaeodactylum tricornutum* inducing a significant increase in the unsaturated fatty acids [21] . Upon completion of elongation, FAs are transported to the cytoplasm to act as substrates of the acyl transferases involved in the TAG synthesis. TAG are neutral lipids formed by the esterification of one molecule of glycerol with three FAs. Because of their energy-rich acyl chains, they are the dominant form of stored energy in microalgae. Cellular stresses, such as nutrient deprivation (carbon dioxide, nitrogen, silica, and phosphorous), temperature fluctuation, or high light exposure trigger their formation [22–28]. It has been demonstrated that lipid biosynthetic pathways are induced under these conditions to potentiate the lipid storage (30–60% of dry cell weight), and this mechanism is thought to play a role in microalgae adaptation and survival [24, 29–39]. It has further been reported that multiple stressors have

Data relating to plant FA and TAG metabolism provided the key to identifying possible molecular targets involved in lipid synthesis and accumulation in microalgae [41]. As shown in **Figure 2**, in plants, the first step of the conventional Kennedy pathway involves the acylation of the glycerol-3-phosphate (G-3-P), catalyzed by the glycerol-3-phosphate acyltransferase (GPAT) to yield lysophosphatidic acid (LPA). GPAT is the rate-limiting step subject to many regulatory controls at the transcriptional and post-transcriptional level and to allosteric mechanisms [42, 43]. Recent studies have revealed the presence of this enzyme in microalgae. In the marine diatom *T. pseudonana*, a membrane-bound GPAT designated TpGPAT was cloned and characterized. The authors observed that G-3-P was the preferred substrate of TpGPAT [44]. A sequence for GPAT with high homology to that of plants was found in *C. reinhardtii, Volvox carteri, Ostreococcus lucimarinus, Ostreococcus tauri, Cyanidioschyzon merolae*, and *P. tricornutum*. As in *T. pseudonana*, G-3-P and fatty acyl molecules are likely to be the enzyme

As described in **Figure 2**, lysophosphatidic acid acyltransferase (LPAAT) participates in the second step of the Kennedy pathway. This enzyme catalyzes the acylation of the LPA to yield phosphatidic acid (PA) [46]. Candidate LPAATs have been found in some algal genomes including that of *H. pluvialis* [47, 48], where it has been shown that LPAAT mRNA is induced under high irradiance stress [47]. In addition, it was recently reported that the expression of *C. reinhardtii* LPAAT (CrLPAAT1) is associated with an increase in lipid synthesis and accumula-

Phosphatidic acid phosphohydrolase (PAP) uses PA as substrate to form diacylglycerol (DAG), a precursor of TAG (**Figure 2**) [49]. In eukaryotes, PAP enzymes are the members of the evolutionarily conserved lipin protein family whose activity is related to TAG storage [50]. In the green microalga *C. reinhardtii*, PAP transcripts (named CrPAP2) are induced under stress conditions. In addition, CrPAP2 silencing slightly lowers the lipid content. Thus, in *C. reinhardtii*, as in other eukaryotes, PAP expression is related to lipid synthesis and accumulation [49].

substrates, as suggested by the residues present in their active sites [45].

no additive effect on lipid accumulation [24, 40].

122 Fatty Acids

tion under nitrogen starvation [48].

**Figure 2.** Simplified overview of the pathways involved in TAG synthesis in plants. Enzymes of the conventional Kennedy pathway involved in TAG synthesis and their subcellular localization in plants. Enzyme abbreviations: glycerol-3-phosphate acyltransferase (GPAT); lysophosphatidic acid acyltransferase (LPAAT); phosphatidic acid phosphohydrolase (PAP); diacylglycerol acyltransferase (DAGAT or DGAT). The same enzymes are involved in TAG synthesis in microalgae, but their intracellular localization has not yet been determined.

The last enzyme of the *de novo* TAG synthesis is acyl-CoA:diacylglycerol acyltransferase (DGAT), which catalyzes the acylation of DAG to yield TAG (**Figure 2**) [51]. This enzyme employs DAG and acyl-CoA as substrates, so the resulting TAG is formed through an acyl-CoA-dependent pathway [46] and is a key target to increase TAG synthesis and storage through genetic manipulation [52, 53]. In higher plants, three different types of DGATs participate in the formation of TAG: DGAT1, DGAT2, or DGAT3 [54]. Sequences for DGAT1 and DGAT2 isoforms were found in several algal strains [55]. Sequences for DGAT2, but not DAGT1, or DGAT3, were identified in the green microalga *O. tauri* [56]. DGAT2 was also found in *T. pseudonana* (TpDGAT2). In addition, the expression of DGAT in a TAG-null yeast mutant restored the synthesis of these neutral lipids [57]. In the oleaginous microalga *C. pyrenoidosa* grown under stress conditions, a high correlation was found between DGAT and TAG accumulation [58]. Also in *N. oceanica* IMET1, another oleaginous microalga, seven putative DGAT genes were up-regulated under nitrogendeficient conditions, when the synthesis of TAG-neutral lipids was significantly increased [59]. In *C. reinhardtii* dgat1 and dgtt1 to dgtt5 genes encode for DGAT1 and DGAT2, respectively [60, 61]. Increased transcript expression of the genes dgat1 and dgtt1 was detected under stress conditions (less sulfur, phosphorous, iron, zinc, or nitrogen). Once more, the evidence suggests that both DGAT1 and DGAT2 could play a role in TAG synthesis as their expression is induced under TAG-accumulating conditions [62, 63]. In support of this hypothesis, overexpression of a DGAT2 isoform in the marine diatom *P. tricornutum* stimulated the synthesis of neutral lipids and their accumulation in lipid droplets [64].

As can be observed, much research has focused on the acyl-CoA-dependent reaction catalyzed by DGAT. However, the relative contribution of DGAT1 and DGAT2 isoenzymes to TAG accumulation appears to be species-dependent, so further studies should be performed to gain insight into this aspect.

TAG can be formed by the acyl-CoA-dependent pathway, detailed previously, or through acyl-CoA-independent reactions. Acyl-CoA-independent formation of TAG is mediated by the activities of two types of enzyme: the phospholipid:diacylglycerol acyltransferases (PDAT), which catalyze the formation of TAG using DAG and phosphatidylcholine (PC); and the DAG:DAG transacylases (DGTA) which utilize two molecules of DAG to form TAG and MAG [54, 65].

In fact, in *N. oceanica* IMET1, it was reported that membrane polar lipids were converted into TAG when the microalgae were grown under nitrogen deficiency [59]. In agreement with this, the gene encoding the acyltransferase PDAT1 was induced under nitrogen starvation in *C. reinhardtii*. Moreover, TAG content in the *C. reinhardtii* PDAT-null mutant was 25% lower than in the parent strain. It would thus appear that PDAT has a relevant role in TAG accumulation, stimulating the transacylation pathway in both strains [62]. Furthermore, in *C. reinhardtii* it was suggested that PDAT functions as a DGTA with acyl hydrolase activity. PDAT might, therefore, mediate membrane polar lipid turnover in a favorable environment whereas under stress conditions it may participate in phospholipid degradation contributing to TAG synthesis [66].

As already mentioned, many aspects of *C. reinhartii* lipid metabolism have already been characterized, making it the microalga of choice for current purposes [23, 67–73]. Nevertheless, *Chlamydomonas* is a non-oleaginous strain [23]. Other microalgal species with greater potential to yield biodiesel and other high-value products should therefore be more thoroughly investigated.

### **3. Transcriptomics, proteomics, and metabolomics**

A better understanding of the mechanisms involved in TAG enrichment under stress conditions will help to maximize microalgae productivity. However, many biochemical approaches for elucidating molecular pathways depend on the availability of genomic sequence data [29]. Transcriptomics, proteomics, and metabolomics, however, are able to provide a detailed description of cell transcripts (RNA), proteins and metabolites, respectively while completely bypassing the requirement of genomic information [74, 75].

Transcriptome analysis helped to identify sequences of the enzymes involved in the biosynthesis and catabolism of FA, TAG, and starch in *D. tertiolecta*, revealing that this strain shares genetic information, at least in terms of the mentioned pathways, with closely related microalgae species such as *V. carteri* and *C. reinhardtii* [76]. The transcriptome of *N. oleoabundans* was also determined. In this case, the authors quantified the differences between nitrogen-replete and nitrogen-limiting culture conditions. Under nitrogen deficiency, *N. oleoabundans* showed higher levels of transcripts of FA and TAG synthesis pathways and inhibition of the FA β-oxidation pathway, compared to nitrogen-replete culture conditions [29]. In agreement with this finding, in *C. vulgaris*, transcriptomic [31] and proteomic [77] studies revealed an induction of the enzymes of the FA and TAG synthesis machinery under lipid enrichment conditions. Also, transcription factors associated with these metabolic pathways were augmented under the stress condition [77].

The transcriptome of *C. reinhardtii* showed that genes involved in FA and TAG metabolic pathways and in membrane remodeling were highly induced under neutral lipid accumulation conditions [78]. In this microalga, proteomic studies revealed an augmented rate of lipid synthesis machinery with a concomitant enhancement in FA and TAG; higher levels of starch than under non-stress conditions were also detected by metabolomic analyses. Metabolic pathways such as nitrogen assimilation, amino acid metabolism, oxidative phosphorylation, glycolysis, TCA cycle, and the Calvin cycle suffered adjustments during *C. reinhardtii* [79, 80].

As in *C. vulgaris*, nutrient-deprivation stress in *C. reinhardtii, D. tertiolecta*, and *N. oleoabundans* induced the expression of genes involved in FA and TAG synthesis pathways in *P. tricornutum* [81], *Chlorella protothecoides* [82], and *Tisochrysis lutea* [83].

In conclusion, these assembled transcriptomes, proteomes, and metabolomes offer valuable approaches for improving microalgal productivity, providing possible targets for molecular engineering that could enhance microalgae-derived products.

### **4. Molecular targets for enhancing lipid biosynthesis**

under TAG-accumulating conditions [62, 63]. In support of this hypothesis, overexpression of a DGAT2 isoform in the marine diatom *P. tricornutum* stimulated the synthesis of neutral lipids

As can be observed, much research has focused on the acyl-CoA-dependent reaction catalyzed by DGAT. However, the relative contribution of DGAT1 and DGAT2 isoenzymes to TAG accumulation appears to be species-dependent, so further studies should be performed

TAG can be formed by the acyl-CoA-dependent pathway, detailed previously, or through acyl-CoA-independent reactions. Acyl-CoA-independent formation of TAG is mediated by the activities of two types of enzyme: the phospholipid:diacylglycerol acyltransferases (PDAT), which catalyze the formation of TAG using DAG and phosphatidylcholine (PC); and the DAG:DAG transacylases (DGTA) which utilize two molecules of DAG to form TAG and MAG [54, 65].

In fact, in *N. oceanica* IMET1, it was reported that membrane polar lipids were converted into TAG when the microalgae were grown under nitrogen deficiency [59]. In agreement with this, the gene encoding the acyltransferase PDAT1 was induced under nitrogen starvation in *C. reinhardtii*. Moreover, TAG content in the *C. reinhardtii* PDAT-null mutant was 25% lower than in the parent strain. It would thus appear that PDAT has a relevant role in TAG accumulation, stimulating the transacylation pathway in both strains [62]. Furthermore, in *C. reinhardtii* it was suggested that PDAT functions as a DGTA with acyl hydrolase activity. PDAT might, therefore, mediate membrane polar lipid turnover in a favorable environment whereas under stress conditions it may participate in phospholipid degradation contribut-

As already mentioned, many aspects of *C. reinhartii* lipid metabolism have already been characterized, making it the microalga of choice for current purposes [23, 67–73]. Nevertheless, *Chlamydomonas* is a non-oleaginous strain [23]. Other microalgal species with greater potential to yield biodiesel and other high-value products should therefore be more thoroughly investigated.

A better understanding of the mechanisms involved in TAG enrichment under stress conditions will help to maximize microalgae productivity. However, many biochemical approaches for elucidating molecular pathways depend on the availability of genomic sequence data [29]. Transcriptomics, proteomics, and metabolomics, however, are able to provide a detailed description of cell transcripts (RNA), proteins and metabolites, respectively while completely

Transcriptome analysis helped to identify sequences of the enzymes involved in the biosynthesis and catabolism of FA, TAG, and starch in *D. tertiolecta*, revealing that this strain shares genetic information, at least in terms of the mentioned pathways, with closely related microalgae species such as *V. carteri* and *C. reinhardtii* [76]. The transcriptome of *N. oleoabundans* was also determined. In this case, the authors quantified the differences between nitrogen-replete and nitrogen-limiting

**3. Transcriptomics, proteomics, and metabolomics**

bypassing the requirement of genomic information [74, 75].

and their accumulation in lipid droplets [64].

to gain insight into this aspect.

124 Fatty Acids

ing to TAG synthesis [66].

Genetic strain modification to improve microalgal productivity and accelerate the industrialization of algal-derived products is a major challenge [84]. Reflecting the fact that enhancement of the FA synthesis pathway had little effect on total lipid content in some plants [85, 86], a growing body of research now focuses on overexpression of the enzymes or heterologous expression of genes involved in the TAG biosynthetic pathway. **Table 1** provides an outline of some of the genetic manipulations performed on several microalgal strains, leading to an improvement in their TAG content.



Enzyme abbreviations: ME, malic enzyme; DGAT, diacylglycerol acyltransferase; G3PDH, glycerol-3-phosphatedehydrogenase; GPAT, glycerol-3-phosphate acyltransferase; LPAAT, lysophosphatidic acid acyltransferase; PAP, phosphatidic acid phosphatase.

**Table 1.** Some of the genetic modifications performed on metabolic pathways related to lipid synthesis in microalgae and their effect on lipid enrichment.

### **5. TAG-accumulation in lipid droplets**

Lipid droplets (LDs) are cell organelles that are currently the subject of in-depth study in various organisms. These lipid globules not only act as a reservoir of cell carbon and energy, they may also have a role in lipid homeostasis, signaling, trafficking, and interorganelle communications [96, 97]. As previously mentioned, under stress conditions microalgae synthesize TAG and store them as cytoplasmic LDs [22–28], which can vary in size, shape, and function depending on the cell type and the environmental conditions (**Figure 3**) [98]. In eukaryotic cells, LD structure consists of a TAG-rich hydrophobic core surrounded by surface polar glycerolipids into which proteins of the perilipin (Plin) (animal cells) or oleosin and caleosin (plants) families are embedded [99–102]. In microalgae, LD structure is conserved from eukaryotes but different LD proteins have been identified. The analysis of *C. reinhardtii* LDs recognized 16 proteins related to lipid metabolism and a major lipid droplet protein (named MLDP) was identified. MLDP silencing increased the size of the LD, without modifying LD TAG content [68]. In the green microalga, *Nannochloropsis* sp., a hydrophobic lipid droplet surface protein, named LDSP, was identified. The expression of LDSP increased concomitantly with TAG content under oil-accumulating conditions [99]. In *H. pluvialis*, seven proteins were found to be

**Figure 3.** Schematic representation of a cytoplasmic lipid droplet (LD) from microalgae.

**5. TAG-accumulation in lipid droplets**

**Enzymes overexpressed or heterologously expressed**

G3PDH, GPAT, DGAT, LPAAT and PAP of *Saccharomyces cerevisiae* and *Yarrowia lipolytica*

LPAAT of *Chlamydomonas* 

DGAT2 of *Nannochloropsis* 

DGAT1 and DGAT2 of *Myrmecia incisa*

DGAT2 of *Neochloris oleoabundans*

DGAT 1 of *Phaeodactylum* 

DGAT 2 of *Phaeodactylum* 

phosphatidic acid phosphatase.

and their effect on lipid enrichment.

*reinhardtii*

126 Fatty Acids

*oceanica*

*tricornutum*

*tricornutum*

Lipid droplets (LDs) are cell organelles that are currently the subject of in-depth study in various organisms. These lipid globules not only act as a reservoir of cell carbon and energy, they may also have a role in lipid homeostasis, signaling, trafficking, and interorganelle communications [96, 97]. As previously mentioned, under stress conditions microalgae synthesize TAG and store them as cytoplasmic LDs [22–28], which can vary in size, shape, and function depending on the cell type and the environmental conditions (**Figure 3**) [98]. In eukaryotic cells, LD structure consists of a TAG-rich hydrophobic core surrounded by surface polar glycerolipids into which proteins of the perilipin (Plin) (animal cells) or oleosin and caleosin (plants) families are embedded [99–102]. In microalgae, LD structure is conserved from eukaryotes but different LD proteins have been identified. The analysis of *C. reinhardtii* LDs recognized 16 proteins related to lipid metabolism and a major lipid droplet protein (named MLDP) was identified. MLDP silencing increased the size of the LD, without modifying LD TAG content [68]. In the green microalga, *Nannochloropsis* sp., a hydrophobic lipid droplet surface protein, named LDSP, was identified. The expression of LDSP increased concomitantly with TAG content under oil-accumulating conditions [99]. In *H. pluvialis*, seven proteins were found to be

**Organism Effect on lipid production** 

**(changes over control** 

Re-stored TAG formation [93]

Re-stored TAG formation [94]

Re-stored TAG synthesis and lipid body formation **References**

[63]

**condition)**

*Chlorella minutissima* 2-fold [90]

*Chlamydomonas reinhardtii* 20% [91]

*Nannochloropsis oceanica* 3.5-fold [92]

*Phaeodactylum tricornutum* 35% [64]

Enzyme abbreviations: ME, malic enzyme; DGAT, diacylglycerol acyltransferase; G3PDH, glycerol-3-phosphatedehydrogenase; GPAT, glycerol-3-phosphate acyltransferase; LPAAT, lysophosphatidic acid acyltransferase; PAP,

**Table 1.** Some of the genetic modifications performed on metabolic pathways related to lipid synthesis in microalgae

*S. cerevisiae* lipid deficient

*S. cerevisiae* DGAT deficient

*S. cerevisiae* DGAT deficient

Various DGAT type 2 *Chlamydomonas reinhardtii* 20–44% [95]

mutant

mutant

mutant

associated with LDs. The most abundant of these, *Haematococcus* Oil Globule Protein (HOGP), was homologous to the MLDP of *C. reinhardtii* and its expression was induced under TAG accumulating conditions [103]. LD-associated proteins may also help in the accumulation of TAG in the green microalga *Myrmecia incisa* [104]. Moreover, LDs from *C. reinhardtii* showed the presence of enzymes involved in TAG synthesis (GPAT, and PDAT) and in sterol synthesis, lipid signaling, and trafficking [69]. Further in-depth research should be able to determine the proteins associated with LDs and their role in TAG metabolism in microalgae.

In the oleaginous diatom *Fistulifera* sp., two proteins located in the oil bodies were also detected in the endoplasmic reticulum (ER), suggesting that oil bodies might originate in the ER [105]. The same authors found a signal sequence typical of ER localization in an LD protein called diatom-oleosome-associated-protein 1 (DOAP1) in *Fistulifera solaris* JPCC DA0580 [106]. Related to these findings, the induction of ER stress leads to LD formation in *C. reinhardtii* and *C. vulgaris* [107]. In addition, LDs from *C. reinhardtii* were associated not only with the ER membrane but also with the outer membrane of the chloroplasts [108]. Available data therefore suggest that in microalgae, cytoplasmic LDs are produced in the ER. However, additional studies are required to arrive at a better understanding of the mechanism of LD formation in the ER, and to determine whether chloroplasts play a role in this process.

### **6. TAG degradation pathways in microalgae**

As previously mentioned, the economic feasibility of using microalgae as a source of FA for biodiesel depends to a great extent on improvements in the production process, one of the most significant challenges being to increase lipid yields. The selection of oleaginous strains and the search for different culture strategies to increase lipid biosynthesis constitute viable approaches; blocking the competing pathways of carbohydrate formation may be another. However, both the approaches give rise to a decrease in strain growth [22]. Lipid catabolism has largely been ignored as a relevant pathway for engineering, despite being a competing pathway to lipid biogenesis [109]. However, lipases were identified in *C. reinhardtii* [66, 72, 73] and *T. pseudonana* [110]. In the case of *C. reinhardtii*, CrLIP1 could restore the lipase activity in a *Saccharomyces cerevisiae* lipase-null strain. In addition, *C. reinhardtii* TAG content decreased with increasing expression of CrLIP1 under stress conditions, hydrolyzing mainly DAG and polar lipids [72]. In agreement with this, a galactoglycerolipid lipase was found in *C. reinhardtii*. The main substrates of the enzyme are galactoglycerolipids and the main products are FAs employed for TAG synthesis [74]. In *C. reinhardtii*, phospholipid:diacylglycerol acyltransferase (PDAT) demonstrated both transacylation and acyl hydrolase activities, and could mediate membrane lipid turnover and TAG synthesis [66]. The activity of a multifunctional lipase/phospholipase/ acyltransferase of *T. pseudonana* lowered lipid content under both normal and stress conditions [110]. A single gene for PDAT was identified in *H. pluvialis*, though no functional analysis was performed for the gene in this strain [47]. Further studies are required to gain insight into the molecular mechanisms involved in TAG degradation, which could be the key to increased lipid yields in microalgae.

### **7. Microalgae-based biorefineries**

In the context of improving the economic feasibility of microalgae-based biodiesel, a closer look should be taken at the large amounts of TAG produced in some oleaginous microalgae alongside high-value products such as carbohydrates (cellulose and starch); proteins and other high-value compounds like pigments, antioxidants (i.e., β-carotene, astaxanthin), and vitamins [2, 3, 8, 9], all of which may have commercial application in different industrial sectors. Some potentially high-value products found in microalgae are described in **Table 2**.



**Table 2.** Recent advances in microalgal-derived high-value products.

### **8. Conclusion**

**6. TAG degradation pathways in microalgae**

yields in microalgae.

128 Fatty Acids

Carbohydrates

Sulfated extracellular polisaccharides

Lipids

**7. Microalgae-based biorefineries**

As previously mentioned, the economic feasibility of using microalgae as a source of FA for biodiesel depends to a great extent on improvements in the production process, one of the most significant challenges being to increase lipid yields. The selection of oleaginous strains and the search for different culture strategies to increase lipid biosynthesis constitute viable approaches; blocking the competing pathways of carbohydrate formation may be another. However, both the approaches give rise to a decrease in strain growth [22]. Lipid catabolism has largely been ignored as a relevant pathway for engineering, despite being a competing pathway to lipid biogenesis [109]. However, lipases were identified in *C. reinhardtii* [66, 72, 73] and *T. pseudonana* [110]. In the case of *C. reinhardtii*, CrLIP1 could restore the lipase activity in a *Saccharomyces cerevisiae* lipase-null strain. In addition, *C. reinhardtii* TAG content decreased with increasing expression of CrLIP1 under stress conditions, hydrolyzing mainly DAG and polar lipids [72]. In agreement with this, a galactoglycerolipid lipase was found in *C. reinhardtii*. The main substrates of the enzyme are galactoglycerolipids and the main products are FAs employed for TAG synthesis [74]. In *C. reinhardtii*, phospholipid:diacylglycerol acyltransferase (PDAT) demonstrated both transacylation and acyl hydrolase activities, and could mediate membrane lipid turnover and TAG synthesis [66]. The activity of a multifunctional lipase/phospholipase/ acyltransferase of *T. pseudonana* lowered lipid content under both normal and stress conditions [110]. A single gene for PDAT was identified in *H. pluvialis*, though no functional analysis was performed for the gene in this strain [47]. Further studies are required to gain insight into the molecular mechanisms involved in TAG degradation, which could be the key to increased lipid

In the context of improving the economic feasibility of microalgae-based biodiesel, a closer look should be taken at the large amounts of TAG produced in some oleaginous microalgae alongside high-value products such as carbohydrates (cellulose and starch); proteins and other high-value compounds like pigments, antioxidants (i.e., β-carotene, astaxanthin), and vitamins [2, 3, 8, 9], all of which may have commercial application in different industrial sectors. Some potentially high-value products found in microalgae are described in **Table 2**.

agronomics

*Graesiella* sp. Pharmaceutics [114]

[111]

**Product Microalgal strain Applications References**

Starch, glucose, cellullose *Chlorella vulgaris FSP-E* Bioethanol production [112, 113]

Exopolysaccharides *Navicula cincta* Pharmaceutics and

Oleaginous microalgae grown under stress conditions can synthesize and accumulate large quantities of FA, mainly in the form of TAG, which can then be converted into biodiesel. Although microalgae constitute a promising source of clean energy, knowledge gaps continue to abound in almost all aspects of FA and TAG metabolism for these microorganisms, including the precise identity of enzymatic machinery, the relative contributions of each enzyme and their precise regulation. Further studies are therefore required to establish the exact metabolic pathways involved in FA and TAG synthesis, accumulation, and degradation in order to develop genetic engineering strategies to obtain microalgal strains with improved capacity to convert their biomass into TAG and other valuable co-products.

### **Acknowledgements**

The authors are grateful for research funds provided by the Consejo Nacional de Investigaciones Científicas y Técnicas de la República Argentina (CONICET); Agencia Nacional de Promoción Científica y Tecnológica, PICTs 2014-0893, 2013-0987, and 2015-0800; and the Secretaría de Ciencia y Tecnología de la Universidad Nacional del Sur, PGIs 24/B226 and 24/B196. Paola Scodelaro Bilbao, Gabriela Salvador and Patricia Leonardi are Research Members of CONICET.

### **Author details**

Paola Scodelaro Bilbao1,2,3\*, Gabriela A. Salvador2,3 and Patricia I. Leonardi1,2

\*Address all correspondence to: pscodela@criba.edu.ar

1 Laboratorio de Estudios Básicos y Biotecnológicos en Algas (LEBBA), Centro de Recursos Naturales Renovables de la Zona Semiárida (CERZOS), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Camino de La Carrindanga, Bahía Blanca, Argentina

2 Universidad Nacional del Sur (UNS), Departamento de Biología, Bioquímica y Farmacia, San Juan, Bahía Blanca, Argentina

3 Instituto de Investigaciones Bioquímicas de Bahía Blanca (INIBIBB), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Camino de La Carrindanga, Bahía Blanca, Argentina

### **References**


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enzyme and their precise regulation. Further studies are therefore required to establish the exact metabolic pathways involved in FA and TAG synthesis, accumulation, and degradation in order to develop genetic engineering strategies to obtain microalgal strains with improved

The authors are grateful for research funds provided by the Consejo Nacional de Investigaciones Científicas y Técnicas de la República Argentina (CONICET); Agencia Nacional de Promoción Científica y Tecnológica, PICTs 2014-0893, 2013-0987, and 2015-0800; and the Secretaría de Ciencia y Tecnología de la Universidad Nacional del Sur, PGIs 24/B226 and 24/B196. Paola Scodelaro Bilbao, Gabriela Salvador and Patricia Leonardi are Research

1 Laboratorio de Estudios Básicos y Biotecnológicos en Algas (LEBBA), Centro de Recursos Naturales Renovables de la Zona Semiárida (CERZOS), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Camino de La Carrindanga, Bahía Blanca, Argentina

2 Universidad Nacional del Sur (UNS), Departamento de Biología, Bioquímica y Farmacia,

3 Instituto de Investigaciones Bioquímicas de Bahía Blanca (INIBIBB), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Camino de La Carrindanga, Bahía Blanca,

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Paola Scodelaro Bilbao1,2,3\*, Gabriela A. Salvador2,3 and Patricia I. Leonardi1,2

\*Address all correspondence to: pscodela@criba.edu.ar

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**Acknowledgements**

130 Fatty Acids

Members of CONICET.

San Juan, Bahía Blanca, Argentina

**Author details**

Argentina

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### **Chapter 8**

## **Fatty Acids in Fish**

Oğuz Taşbozan and Mahmut Ali Gökçe

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/68048

#### **Abstract**

The human body cannot synthesize certain fatty acids: these essential fatty acids must be consumed in the diet. Fish and other aquatic foods are known to be the main sources of polyunsaturated fatty acids (PUFA); therefore, humans obtain most of their eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) by consuming fish, aquatic invertebrates, and algae. The increasing demand for fish and the stabilization of marine fish and freshwater landings have contributed to a widening gap between demand and supply for fish and fish products. This leads to a necessity to improve aquaculture production. Fish are the main contributors of n‐3 PUFA in the human diet, although there are some interspecific and intraspecific differences in fatty acid profiles. The fatty acid composition of fish differs depending on a variety of factors, including species, diet, as well as environmental factors such as salinity, temperature, season, geographical location, and whether the fish are farmed or wild. In this chapter, information will be provided on fish fatty acids based on their ecology, feeding habits, lipid contents, and environmental conditions where they are harvested.

**Keywords:** marine fish, freshwater fish, EPA, DHA, PUFA, HUFA, n3/n6

### **1. Introduction**

Many studies have investigated the effects of lipids and fatty acids in human nutrition on health. This has resulted in an increasing consumer interest and a tendency to consume healthy foods.

Among the fatty acids, highly unsaturated n‐3 fatty acids (n‐3 HUFA) or long‐chain n‐3 poly unsaturated fatty acids (LC n‐3 PUFA), particularly 20:5 n‐3 (eicosapentaenoic acid [EPA]) and 22:6 n‐3 (docosahexaenoic acid [DHA]) affect human health, early development, and the prevention of some diseases; therefore, dieticians increasingly recommend consuming foods

© 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

containing these fatty acids [1]. The recommended n‐6/n‐3 fatty acid ratio in human nutrition is 5:1, but this ratio (n‐6/n‐3) varies between 7:1 and 20:1 in the diets of most West Europeans and North Americans [1, 2]. The n‐3/n‐6 fatty acid ratio recommended by the World Health Organization is 1:1 or above [3]; hence, fish consumption should be increased or foods rich in n‐3 fatty acids should be consumed for proper nutrition and disease prevention.

Fish are the most important sources of these fatty acids; fatty fish, such as sardines, mackerel, anchovies, and some salmon species, are rich in EPA and DHA. In these fish, the ratio of n‐3 fatty acid to n‐6 fatty acid approaches 7. Fish cannot synthesize these fatty acids; they obtain them from food they consume (algae and planktons) [4].

However, lipid composition and thus fatty acid composition in fish differ depending on various factors: usually, their aquatic environment (marine water, freshwater, and cold or warm water) and the biological, physical, and chemical properties of that environment. Also, seasonal changes, migration, sexual maturity and spawning period, species, feeding habits, and whether reared in aquaculture or grown in natural habitats affect the lipid/fatty acid composition [5].

Therefore, detailed information on the changes in lipid and fatty acid compositions among fish and the importance of fish consumption in human health are provided in this chapter by examining these subjects.

### **2. The importance of fatty acids and fish consume**

The interest in fat, which holds an important place in human nutrition, has increased with the recently increasing interest in, and awareness of, human health. Fats are important components of hormones, cell membranes, and signaling molecules, as well as being important energy sources. Fat ingested into the body is first stored in the liver, hypodermic connective tissues, mesentery, and muscles and used when necessary [6].

Fatty acids have a methyl group on one end and have long hydrocarbon chains carrying a carboxyl group on the other end. Fatty acid molecules are classified based on the presence and number of double bonds: saturated fatty acids have no double bond, and monounsaturated fatty acids have a single double bond; polyunsaturated fatty acids (PUFA) have two or more double bonds. The number and position of the double bonds determine the physical properties and functional characteristics of fatty acids. The human body can produce some of these fatty acids, but others, some of which also contain n‐3 and n‐6 PUFA, cannot be produced by the body. These essential fatty acids (EFAs) need to be obtained through food intake. In current human diets, n‐6 fatty acid and, especially, linoleic acid (LA) sources are soya and maize oil, and arachidonic acid (AA); the main n‐3 alpha‐linolenic acid (ALA) source is meat. Linoleic acid (LA), an n‐6 fatty acid, can be converted to fatty acids with longer chains, and n‐3 ALA can be converted to eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA); these conversion rates vary between 1 and 10%. Even though EPA and DHA, with 20–22 long chain n‐3 fatty acids, have a critical role in human health, their consumption is relatively low, stemming from the deficiency in consumption of fish and fish products in developed countries [7–9].

Many recent studies have shown the importance of and necessity for n‐3 fatty acids in human development and health. Some studies show that they have a positive effect on maternal and fetal health during pregnancy and on newborn and childhood health. These studies emphasize an important role for fatty acids in prevention of hormone‐related cancers and important functions in the prevention of cardiovascular diseases. These fatty acids also are purported to relieve dementia, hyperactivity, and some psychiatric disorders [10].

Many studies have carefully evaluated the effects of the lack of fatty acids in the diet following the onset of pregnancy on the prenatal and on postnatal development of newborns and children [6, 10–12]. One study investigated the effect of the lack of n‐6 and n‐3 long chain PUFA on children with attention‐deficit hyperactivity [13]. Moreover, several studies have evaluated the relationship between n‐3 PUFA deficiency and depression and mood disorders [14–16]. Various studies have reported the cardio‐protective effect of n‐3 PUFA (EPA and DHA) supplementation and recommend 1 g EPA intake per day to prevent coronary heart disease [17]. Although their anti‐cancer roles have yet to be proven, many studies have shown that n‐3 and n‐6 PUFA positively affected the prevention of development of different types of tumors. The n‐3 PUFA has been reported to alter cell growth by modulating cell replication, interfering with the components of the cell cycle, or by increasing cell death through necrosis or apoptosis [18, 19]. Other studies on the same subject have focused on breast cancer, colon and colorectal cancers, prostate cancer, lung cancer, and neuroblastoma [20].

Fish consumption has an important role in n‐3 PUFA (EPA and DHA) intake. Although fatty acids of the n‐3 group that cannot be synthesized in the body vary in different species and individuals, fish contain significant amounts of n‐3 fatty acids. The nutritional contents and fatty acid compositions of fish differ, depending on various factors. For example, fatty fish, such as herring and mackerel have 400 mg of PUFA per 15 g; thus, weekly consumption of 300 g fatty fish or a daily 200 mg EPA and DHA intake is reported to be sufficient [21]. Furthermore, the n‐3/n‐6 ratio is reported to be a good index for comparison of the relative nutritional value of fish oils [22]. Although there is no specific recommendation for the n‐3/n‐6 ratio, evidence found in wild animals and estimated food intake during human evolution indicates a dietary ratio of 1:1 [3].

### **3. How the fatty acid profiles vary in fish?**

containing these fatty acids [1]. The recommended n‐6/n‐3 fatty acid ratio in human nutrition is 5:1, but this ratio (n‐6/n‐3) varies between 7:1 and 20:1 in the diets of most West Europeans and North Americans [1, 2]. The n‐3/n‐6 fatty acid ratio recommended by the World Health Organization is 1:1 or above [3]; hence, fish consumption should be increased or foods rich in

Fish are the most important sources of these fatty acids; fatty fish, such as sardines, mackerel, anchovies, and some salmon species, are rich in EPA and DHA. In these fish, the ratio of n‐3 fatty acid to n‐6 fatty acid approaches 7. Fish cannot synthesize these fatty acids; they obtain

However, lipid composition and thus fatty acid composition in fish differ depending on various factors: usually, their aquatic environment (marine water, freshwater, and cold or warm water) and the biological, physical, and chemical properties of that environment. Also, seasonal changes, migration, sexual maturity and spawning period, species, feeding habits, and whether reared in aquaculture or grown in natural habitats affect the lipid/fatty acid composition [5].

Therefore, detailed information on the changes in lipid and fatty acid compositions among fish and the importance of fish consumption in human health are provided in this chapter by

The interest in fat, which holds an important place in human nutrition, has increased with the recently increasing interest in, and awareness of, human health. Fats are important components of hormones, cell membranes, and signaling molecules, as well as being important energy sources. Fat ingested into the body is first stored in the liver, hypodermic connective

Fatty acids have a methyl group on one end and have long hydrocarbon chains carrying a carboxyl group on the other end. Fatty acid molecules are classified based on the presence and number of double bonds: saturated fatty acids have no double bond, and monounsaturated fatty acids have a single double bond; polyunsaturated fatty acids (PUFA) have two or more double bonds. The number and position of the double bonds determine the physical properties and functional characteristics of fatty acids. The human body can produce some of these fatty acids, but others, some of which also contain n‐3 and n‐6 PUFA, cannot be produced by the body. These essential fatty acids (EFAs) need to be obtained through food intake. In current human diets, n‐6 fatty acid and, especially, linoleic acid (LA) sources are soya and maize oil, and arachidonic acid (AA); the main n‐3 alpha‐linolenic acid (ALA) source is meat. Linoleic acid (LA), an n‐6 fatty acid, can be converted to fatty acids with longer chains, and n‐3 ALA can be converted to eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA); these conversion rates vary between 1 and 10%. Even though EPA and DHA, with 20–22 long chain n‐3 fatty acids, have a critical role in human health, their consumption is relatively low, stemming from the deficiency in consumption of fish and fish products in developed countries [7–9].

n‐3 fatty acids should be consumed for proper nutrition and disease prevention.

them from food they consume (algae and planktons) [4].

**2. The importance of fatty acids and fish consume**

tissues, mesentery, and muscles and used when necessary [6].

examining these subjects.

144 Fatty Acids

Among vertebrates, fish have the highest species diversity and, as stated previously, the nutritional content and fatty acid compositions of fish vary. The most important causes of the variation in the fatty acid profiles of fish are the differences among species. Moreover, living in aquatic environments with different ecological conditions is an important source of variation in nutritional components. The fishing season, size, and reproduction status of the individuals from the same species living in a certain region also affect this variation. Moreover, the aquaculture conditions and feeds used in fish aquaculture also cause variations in the fatty acid compositions of fish that were supplied to the market using aquaculture. These factors are elaborated in the sub‐sections that follow.

#### **3.1. Fish bioecology**

Fish are divided into two groups based on their habitat: marine fish and freshwater fish. Water temperature and salinity are the most important environmental factors; thus, fishes are first studied based on water temperature and then divided into two groups, the warm‐water fish group and cold‐water fish group. The optimal temperatures for warm‐water species are around 25–30°C, whereas cold‐water species prefer temperatures below 20°C. In addition to this classification, both cold‐water and warm‐water fish are further classified as freshwater and marine fish. Moreover, some fish species migrate from seas to fresh waters or from fresh waters to seas. These fish, such as salmonids and eels, are termed diadromous species [23]. The nutritional compositions of fish (especially lipids and fatty acids) substantially vary due to the differences in their habitats.

In general, freshwater fish require either linoleic acid (18:2 n‐6), linolenic acid (18:3 n‐3) or both, whereas marine fish require EPA (20:5 n‐3) and/or DHA (22:6 n‐3) [24].

The studies on the essential fatty acid (EFA) requirement of marine fish showed that their n‐3 EFA requirement can be met by EPA (20:5 n‐3) and DHA (22:6 n‐3). These two fatty acids are termed n‐3 HUFA (highly unsaturated fatty acid) or LC (long chain) n‐3 PUFA (poly unsaturated fatty acid). The EPA and DHA requirements of fish respond to the n‐3 HUFA rich nutrients in marine environments because primary food sources, such as marine algae and planktons, and also other food sources, are known to be rich in EPA and DHA and contain lower levels of linoleic acid and linolenic acid [25, 26].

Studies on freshwater fish have shown that their n‐3 EFA requirements are mostly focused on linolenic acid (18:3n‐3). The fact that EPA and DHA in vertebrates are biologically active forms of n‐3 EFA led to the conclusion that many freshwater fish can convert 18:3 n‐3 fatty acids to EPA and DHA. This is less straightforward in marine fish. When compared to marine microalgae, freshwater microalgae contain higher levels of 18:3 n‐3 fatty acids than EPA and DHA. Moreover, even though there is not a great amount of 18:2 n‐6 fatty acid in marine microalgae, freshwater microalgae contain plenty of this fatty acid. This explains why freshwater fish require higher amounts of linoleic and linoleic acid as compared to marine fish [25, 26]. On the other hand, only one species among the freshwater fish (*Esox lucius*) has the ability to convert 18:3 n‐3 fatty acid to EPA and DHA. Because this fish species is an extreme carnivore consuming smaller fish, it does not have high amounts of 18:3 n‐3 and 18:2 n‐6 fatty acids [27].

The nutritional compositions of the foods in the natural environment of marine and freshwater fish necessitate providing farmed fish with food sources that meet the requirements of their species. The essential fatty acid amount required in the feeds of commercial freshwater and marine fish (preadult or older juvenile) is determined in terms of dry diet%. For example, the 18:3 n‐3 fatty acid requirement of trout is 0.7–1.0, and the n‐3 HUFA requirement of trout is 0.4–0.5. The 18:2 n‐6 fatty acid requirement of common carp is 1.0, whereas their 18:3 n‐3 fatty acid requirement ranges from 0.5 to 1.0. The 18:2 n‐6 fatty acid requirements of *Tilapia zilli* and *Oreochromis niloticus* are 1.0 and 0.5, respectively. Channel catfish require 18:3 n‐3 fatty acid between 1.0 and 2.0, and n‐3 HUFA between 0.5 and 0.75. Among the marine fish species, turbot require n‐3 HUFA at a ratio of 0.8, and red sea bream (*Pagrus major*) require n‐3 HUFA at a ratio of 0.5. The EPA and DHA requirements of red sea bream are 1.0 and 0.5, respectively. The n‐3 HUFA requirement of gilthead sea bream (*Sparus aurata*) is between 0.9 and 1.9, whereas the n‐3 HUFA requirement of sea bass (*Dicentrarchus labrax*), another important marine species, is 1.0 [25, 26].

**3.1. Fish bioecology**

146 Fatty Acids

to the differences in their habitats.

and 18:2 n‐6 fatty acids [27].

Fish are divided into two groups based on their habitat: marine fish and freshwater fish. Water temperature and salinity are the most important environmental factors; thus, fishes are first studied based on water temperature and then divided into two groups, the warm‐water fish group and cold‐water fish group. The optimal temperatures for warm‐water species are

this classification, both cold‐water and warm‐water fish are further classified as freshwater and marine fish. Moreover, some fish species migrate from seas to fresh waters or from fresh waters to seas. These fish, such as salmonids and eels, are termed diadromous species [23]. The nutritional compositions of fish (especially lipids and fatty acids) substantially vary due

In general, freshwater fish require either linoleic acid (18:2 n‐6), linolenic acid (18:3 n‐3) or

The studies on the essential fatty acid (EFA) requirement of marine fish showed that their n‐3 EFA requirement can be met by EPA (20:5 n‐3) and DHA (22:6 n‐3). These two fatty acids are termed n‐3 HUFA (highly unsaturated fatty acid) or LC (long chain) n‐3 PUFA (poly unsaturated fatty acid). The EPA and DHA requirements of fish respond to the n‐3 HUFA rich nutrients in marine environments because primary food sources, such as marine algae and planktons, and also other food sources, are known to be rich in EPA and DHA and contain

Studies on freshwater fish have shown that their n‐3 EFA requirements are mostly focused on linolenic acid (18:3n‐3). The fact that EPA and DHA in vertebrates are biologically active forms of n‐3 EFA led to the conclusion that many freshwater fish can convert 18:3 n‐3 fatty acids to EPA and DHA. This is less straightforward in marine fish. When compared to marine microalgae, freshwater microalgae contain higher levels of 18:3 n‐3 fatty acids than EPA and DHA. Moreover, even though there is not a great amount of 18:2 n‐6 fatty acid in marine microalgae, freshwater microalgae contain plenty of this fatty acid. This explains why freshwater fish require higher amounts of linoleic and linoleic acid as compared to marine fish [25, 26]. On the other hand, only one species among the freshwater fish (*Esox lucius*) has the ability to convert 18:3 n‐3 fatty acid to EPA and DHA. Because this fish species is an extreme carnivore consuming smaller fish, it does not have high amounts of 18:3 n‐3

The nutritional compositions of the foods in the natural environment of marine and freshwater fish necessitate providing farmed fish with food sources that meet the requirements of their species. The essential fatty acid amount required in the feeds of commercial freshwater and marine fish (preadult or older juvenile) is determined in terms of dry diet%. For example, the 18:3 n‐3 fatty acid requirement of trout is 0.7–1.0, and the n‐3 HUFA requirement of trout is 0.4–0.5. The 18:2 n‐6 fatty acid requirement of common carp is 1.0, whereas their 18:3 n‐3 fatty acid requirement ranges from 0.5 to 1.0. The 18:2 n‐6 fatty acid requirements of *Tilapia zilli* and *Oreochromis niloticus* are 1.0 and 0.5, respectively. Channel catfish require 18:3 n‐3 fatty acid between 1.0 and 2.0, and n‐3 HUFA between 0.5 and 0.75. Among the marine fish

In addition to

around 25–30°C, whereas cold‐water species prefer temperatures below 20°C.

both, whereas marine fish require EPA (20:5 n‐3) and/or DHA (22:6 n‐3) [24].

lower levels of linoleic acid and linolenic acid [25, 26].

Many studies have focused on determining the lipid and fatty acid compositions of marine and freshwater fish (both cold water and warm water). The goal of these studies was both to find the differences among fatty acid compositions of fish from different aquatic environments and to evaluate these fatty acids in terms of human health.

The results obtained in a study from Turkey on the fatty acid compositions of eight different marine fish species that were either farmed or caught in their natural habitats (waker, tub gurnard, whiting, mackerel, blue fish, sea bream, sea bass, and marbled spinefoot) and six different freshwater fish species caught in their natural habitats offer an insight into this subject. The PUFA ratios of the marine species were between 25.2 and 48.2%, whereas they were between 23.2 and 43.8% in freshwater species. The EPA and DHA values of marine species were 4.23–7.02% and 11.7–36.01%, respectively. The EPA and DHA values of freshwater species were between 2.10 and 13.8%, and between 6.72 and 24.8%, respectively. Among marine fish, at 22.6%, the lowest n‐3 PUFA ratio was found in waker and, at 44.2%, the highest ratio was found in blue fish. Among the freshwater fish, North African catfish had the lowest n‐3 PUFA (11.05%) value, whereas at 28.4%, zander had the highest value. In addition, the n‐6 PUFA ratios in the marine fish were between 0.43 and 14.4% and between 5.27 and 16.8% in the freshwater fish [28]. The researchers reported that n‐6/n‐3 ratios in both the freshwater fish and marine fish were below the ratio recommended by the UK Department of Health (4.0 at maximum) [29].

The results obtained in a study on 34 different marine fish species from the Mediterranean Sea showed that the fatty acid levels of all fish were at the desired levels for human health and quality food consumption. The EPA and DHA values of fish were between 1.94 and 10.0%, and 3.31 and 31.03%, respectively. The n‐3 PUFA levels were between 12.66 (annular sea bream) and 36.54% (European hake), whereas oceanic puffer had the lowest n‐6 PUFA level at 1.24% and flathead mullet had the highest n‐6 PUFA level at 12.76%; the n‐6/n‐3 ratio varied between 0.04 and 0.91 [30].

A study was carried out with marine and freshwater fish from Malaysia found EPA ratios in freshwater fish between 0.63 and 1.41%, whereas they were between 4.68 and 10.62% in marine fish. The DHA levels were between 0.14 and 0.25%, and 2.50 and 10.05% in freshwater fish and marine fish, respectively. Moreover, that study reported that the n‐3/n‐6 ratios in all marine species were above 1, whereas the highest level reached was 0.73 in freshwater fish [31]. The World Health Organization recommended that the n‐3/n‐6 ratio should be at least 1 [3].

Overall, the studies reported that marine species and species that show carnivorous feeding habits and species living in cold water contained high amounts of EPA and DHA and therefore can be used as an important source of food for human health.

#### **3.2. Feeding habits**

Aquatic animals (organisms) have environmental and biological characteristics. The most important biological characteristics are feeding habits. Fish are classified as carnivorous, herbivorous, omnivorous, and detritivorous (detritivore, detrivore, or detritus feeder) based on their usual food source preferences in their natural habitats [32]. Moreover, each class is further classified based on their food source preferences as euryphagous (feed on a great variety of foods), stenophagous (feed on a limited variety of foods), or monophagous (feed on only one type of food) [33]. The detritivorous species is *Cirrhinus molitorella*, known as mud carp, and does not have much commercial value.

The most frequently consumed and farmed fish species worldwide have carnivorous, herbivorous, and omnivorous feeding habits; thus, these fish are rich in nutrients and popular among consumers. The fish species that are widely farmed are: euryphagous carnivores, such as salmon, basses, breams, halibut, flounders, and groupers; euryphagous herbivores, such as some carp and tilapia species, milkfish; or euryphagous omnivores, such as common carp, channel catfish, grey mullet, and eels.

The anatomic structures of fish digestion systems differ depending on their feeding habits. Carnivores have shorter intestines and larger stomachs as compared with the other groups, and their stomachs are usually tube‐shaped [34]. The digestion ratio of food is higher in carnivorous fish when compared with other groups because semi‐digested foods are stored in the chyme, that is, in the stomach, for shorter periods [35].

Common carp, one of the omnivorous species, does not have a stomach because it tends to consume herbal foods; however, some omnivorous species have pouch‐shaped stomachs that are smaller than those of the carnivorous species. Moreover, their intestinal structure is more developed and longer. Herbivorous species do not have stomachs and have the longest and most complex intestinal structure because they consume only herbal food sources [34, 36].

The energy requirements of fish differ depending on their feeding habits; therefore, lipid digestion and requirement for lipids, the most important energy source, vary among the fish species. In addition to fish species and feeding habits, some other factors also affect lipid digestion. The age of the fish is the most important factor in lipid digestion [37–39]. The ability of young fish and, especially, fish at the larval stage to digest foods containing high amounts of lipid and lipids in feeds is markedly insufficient [36, 37, 39, 40]. Temperature also affects lipid digestion: warm‐water fish species have the greatest ability to digest lipids [41, 42].

In general, carnivorous species can better digest the lipids in high‐fat nutrients in their natural habitat—or pellet feeds under farming conditions [43–46]. Their ability to better digest lipids is attributable to their genetic potential to store lipids [47]. In contrast, fish species that have herbivorous and omnivorous feeding habits have a lower capacity to digest nutrients or feeds with high lipid content [48]. Although the ability to digest lipids is affected by many factors, the superior lipid digestion ability of carnivorous species is attributable to their more specific and higher lipase activity when compared with herbivorous and omnivorous species [49].

Based on the results obtained in relevant studies, the fatty acid requirements and compositions of fish were divided into three groups. The first group is the salmon and rainbow trout group, which are freshwater and anadromous carnivore species from the Salmonidae family; the second group is the carnivorous marine fish group which contains sea bass and sea bream; the third group is the temperate‐climate fish group, that mostly have herbivorous and omnivorous feeding habits (tilapia, carp, eel, among others) [50].

For the fish in the first group, α‐linolenic acid (18:3 n‐3) is the main fatty acid that must be in their feeds, especially under farming conditions. Certain levels of EPA and DHA can only be synthesized from linolenic acid by elongation if there is sufficient α‐linolenic acid and insufficient of EPA and DHA in feeds. This does not imply that EPA and DHA are unimportant for trout; on the contrary, trout require these two fatty acids in high amounts but can partially met their requirements with linolenic acid when EPA and DHA are insufficient; however, in some cases, the synthesized amount may itself be insufficient [46, 50].

The most important fatty acids for carnivorous marine fish, especially for sea bass and sea bream, are EPA and DHA. Their ability to synthesize EPA and DHA using other fatty acids is inferior to that of the fish of the Salmonidae group; therefore, they tend to feed on nutrients rich in EPA and DHA, either in their natural habitats or under farming conditions [50].

Linoleic acid (18:2n‐6) is known to be the most important fatty acid requirement of species that have herbivorous and omnivorous feeding habits. These fish species, along with linoleic acid, require linolenic acid (18:3n‐3) and arachidonic acid (20:4n‐6) [50].

**Table 1** shows the fatty acid requirements of some carnivorous and herbivorous fish species [50].

### **3.3. Lipid contents**

**3.2. Feeding habits**

148 Fatty Acids

and does not have much commercial value.

channel catfish, grey mullet, and eels.

[34, 36].

the chyme, that is, in the stomach, for shorter periods [35].

Aquatic animals (organisms) have environmental and biological characteristics. The most important biological characteristics are feeding habits. Fish are classified as carnivorous, herbivorous, omnivorous, and detritivorous (detritivore, detrivore, or detritus feeder) based on their usual food source preferences in their natural habitats [32]. Moreover, each class is further classified based on their food source preferences as euryphagous (feed on a great variety of foods), stenophagous (feed on a limited variety of foods), or monophagous (feed on only one type of food) [33]. The detritivorous species is *Cirrhinus molitorella*, known as mud carp,

The most frequently consumed and farmed fish species worldwide have carnivorous, herbivorous, and omnivorous feeding habits; thus, these fish are rich in nutrients and popular among consumers. The fish species that are widely farmed are: euryphagous carnivores, such as salmon, basses, breams, halibut, flounders, and groupers; euryphagous herbivores, such as some carp and tilapia species, milkfish; or euryphagous omnivores, such as common carp,

The anatomic structures of fish digestion systems differ depending on their feeding habits. Carnivores have shorter intestines and larger stomachs as compared with the other groups, and their stomachs are usually tube‐shaped [34]. The digestion ratio of food is higher in carnivorous fish when compared with other groups because semi‐digested foods are stored in

Common carp, one of the omnivorous species, does not have a stomach because it tends to consume herbal foods; however, some omnivorous species have pouch‐shaped stomachs that are smaller than those of the carnivorous species. Moreover, their intestinal structure is more developed and longer. Herbivorous species do not have stomachs and have the longest and most complex intestinal structure because they consume only herbal food sources

The energy requirements of fish differ depending on their feeding habits; therefore, lipid digestion and requirement for lipids, the most important energy source, vary among the fish species. In addition to fish species and feeding habits, some other factors also affect lipid digestion. The age of the fish is the most important factor in lipid digestion [37–39]. The ability of young fish and, especially, fish at the larval stage to digest foods containing high amounts of lipid and lipids in feeds is markedly insufficient [36, 37, 39, 40]. Temperature also affects lipid digestion: warm‐water fish species have the greatest ability to digest lipids [41, 42].

In general, carnivorous species can better digest the lipids in high‐fat nutrients in their natural habitat—or pellet feeds under farming conditions [43–46]. Their ability to better digest lipids is attributable to their genetic potential to store lipids [47]. In contrast, fish species that have herbivorous and omnivorous feeding habits have a lower capacity to digest nutrients or feeds with high lipid content [48]. Although the ability to digest lipids is affected by many factors, the superior lipid digestion ability of carnivorous species is attributable to their more specific and higher lipase activity when compared with herbivorous and omnivorous species [49].

There are three different lipid compositions of fish: lean fish (less than 5% fat), mid‐fat fish (5–10% fat), and fatty fish (10–25% fat). Although the lipid contents of fish depend on many factors, they are generally divided into three groups based on their composition: lean, mid‐fat or medium fat, and fatty fish. This classification and the lipid levels in fish have been reported somewhat inconsistently in publications and research. For example, in one paper, fish were divided into four groups based on their lipid levels and lean fish were evaluated under two categories. These groups were:

Very low fat (less than 2%): cod, haddock, flounder/sole, and tuna

Low fat (2–5%): tilapia, halibut, ocean perch, and salmon (chum, pink)

Medium fat (5–10%): bluefish, catfish, rainbow trout, and sword fish

High fat (10% or more): herring, mackerel, sardines, and salmon (Atlantic, sockeye, coho, and chinook) [51].

In another study, fish were separated into three different classes. Fish having lipid levels below 2% were regarded as lean fish; fish having lipid levels between 2 and 8% were regarded


**Table 1.** Fatty acid (in dry feed%) requirements of some carnivorous, omnivorous, and herbivorous fish species [50].

as mid‐fat fish; fish having lipid levels above 8% were regarded as fatty fish. Cod fish was given as the best example of lean fish, and some salmon species, herring and mackerel, were placed in the fatty fish group. Another important issue, which should not be overlooked, is that the lipid content in fish can vary significantly. In wild fish, seasonal changes, sexual maturity, reproduction period, and the nutrients they consume; in farmed fish, the feed content and quality directly affect the lipid content [52].

The lipid ratio in lean or fatty fish usually depends on how and where the lipids are stored. Cod fish are known to be lean fish; they do not store lipids in their muscle tissues (fillet) but store them only in the liver, whereas salmon and trout species store lipids in their muscle tissues and the surrounding organs and do not store lipids in their liver [53].

**Table 2** shows the nutritional composition in lean, mid‐fat, and fatty fish [52].

The lipids in fish vary with body composition. In general, the differences in lipid compositions of certain fish species depend on the spawning period and seasonal changes. For instance, seasonal changes were reported to significantly affect the lipid compositions of herring (*Clupea harengus*) and mackerel (*Scomber scombrus*). In herring, the lowest lipid level was around 5% and observed in April; the highest lipid level was above 25% and was observed in July. In fillets of mackerel, the lowest lipid level was 5% and observed between June and July, whereas the lipid level was above 20% during September–January and approached 30% in December [54].

Because these fish are rich in n‐3 fatty acids, the consumption of fatty fish is also important for human health, both EPA and DHA‐rich species including salmon, herring, mackerel, anchovies, and sardines. In these fish, the n‐3 fatty acid contents are sevenfold or more higher than their n‐6 fatty acid contents [55].


**Table 2.** Nutritional content per 100 g in different fish species.

as mid‐fat fish; fish having lipid levels above 8% were regarded as fatty fish. Cod fish was given as the best example of lean fish, and some salmon species, herring and mackerel, were placed in the fatty fish group. Another important issue, which should not be overlooked, is that the lipid content in fish can vary significantly. In wild fish, seasonal changes, sexual maturity, reproduction period, and the nutrients they consume; in farmed fish, the feed con-

**Table 1.** Fatty acid (in dry feed%) requirements of some carnivorous, omnivorous, and herbivorous fish species [50].

**Species Requirements of fatty acids (in dry feed%)**

Linolenic 0.8% EPA + DHA 0.4–0.5%

EPA + DHA 1.9%, EPA:DAHA = 0.5

Rainbow trout (*Oncorhynchus mykiss*) Linolenic 1%

Sea bass (*Dicentrarchus labrax*) EPA + DHA 1%

Sea bream (*Sparus aurata*) EPA + DHA 1%, EPA:DAHA = 1

Common carp (*Cyprinus carpio*) Linoleic 1%; linolenic 0.5–1% Japanese eel (*Anguilla japonicus*) Linoleic 0.5%; linolenic 0.5%

Grass carp (*Ctenopharyngodon idella*) Linoleic 1%; linolenic 0.5% Tilapia (*Tilapia zilli*) Linoleic 1%; arachidonic 1%

Tilapia (*Oreochromis niloticus*) Linoleic 0.5%

The lipid ratio in lean or fatty fish usually depends on how and where the lipids are stored. Cod fish are known to be lean fish; they do not store lipids in their muscle tissues (fillet) but store them only in the liver, whereas salmon and trout species store lipids in their muscle tis-

The lipids in fish vary with body composition. In general, the differences in lipid compositions of certain fish species depend on the spawning period and seasonal changes. For instance, seasonal changes were reported to significantly affect the lipid compositions of herring (*Clupea harengus*) and mackerel (*Scomber scombrus*). In herring, the lowest lipid level was around 5% and observed in April; the highest lipid level was above 25% and was observed in July. In fillets of mackerel, the lowest lipid level was 5% and observed between June and July, whereas the lipid level was above 20% during September–January and approached 30% in December [54]. Because these fish are rich in n‐3 fatty acids, the consumption of fatty fish is also important for human health, both EPA and DHA‐rich species including salmon, herring, mackerel, anchovies, and sardines. In these fish, the n‐3 fatty acid contents are sevenfold or more higher than

sues and the surrounding organs and do not store lipids in their liver [53].

**Table 2** shows the nutritional composition in lean, mid‐fat, and fatty fish [52].

tent and quality directly affect the lipid content [52].

**1. Carnivores**

150 Fatty Acids

**2. Omnivores**

**3. Herbivores**

their n‐6 fatty acid contents [55].

The lipid, EPA, and DHA levels in fillets determined in a study that investigated the lipid levels in fish are given in **Table 3** [52].

In India, researchers examined the EPA, DHA, and fatty acid compositions of 34 fish, 3 prawns, and 2 mussels with different lipid contents (lean, mid‐fat, and fatty) and, again, found that EPA and DHA levels were high in the fatty fish group. Among these fish, 12 were obtained from marine water, 3 were obtained from brackish water, 14 were obtained from freshwater, and 5 were obtained from cold water. *Sardinella longiceps* (marine water) and *Tenualosa ilisha* (freshwater) had the highest lipid contents (9.2 and 10.5%, respectively). It was observed that the fatty acid compositions of both fatty fish were especially rich in HUFA, EPA, DHA, and n‐3 fatty acids. *Sardinella longiceps* had 12.3% EPA, 6.9% DHA, and 21.4% ∑ n‐3 fatty acid. The fatty acid composition of *Tenualosa ilisha,* a freshwater species, contained 2.9% EPA, 8.9% DHA, and 14.2% ∑ n‐3 fatty acid [56].

Although the lipid level of *Catla catla* fish, which is a freshwater species having low lipid levels (2–4%) and was obtained from farms, was low in the body composition, it had high levels of EPA, DHA and ∑ n‐3 fatty acid. Again, although *Rastrelliger kanagurta* and *Stolephorus waitei*


**Table 3.** Fat, EPA, and DHA content in selected fish species.

(lean fish; less than 2% fat) from marine water had low lipid content, they were rich in EPA, DHA, and ∑ n‐3 fatty acid relative to the other fish in the same group [56].

#### **3.4. Wild or farmed fish**

There are significant differences in nutritional compositions of farmed fish and wild fish. Many recent studies have focused on this issue and have tried to determine to what degree the nutritional composition of fish affects human health and has nutritional benefits [57–64]. The nutritional quality of farmed fish has improved in the recent years thanks to environmentally friendly and advanced aquaculture techniques. In addition, the advancing feed sector now can offer the most suitable and best quality feeds.

In its early years, aquaculture was carried out in small areas using artificial feeds and simple techniques, and the only way to handle disease factors was to use antibiotics. However, more recently, aquaculture has improved significantly and has begun to yield quality products that are both environmentally friendly and beneficial to human health. Many studies have shown that the nutritional, and especially, the lipid compositions in farmed fish are more consistent than in wild fish; therefore, they are richer in n‐3 fatty acids [59, 63, 65].

A study that compared individual farmed fish with individual wild fish using the sharp snout sea bream (*Diplodus puntazzo*). In the farmed fish, EPA, DHA, ∑PUFA, ∑n‐3 fatty acid levels, and the n‐3/n‐6 ratio were 4.23 (g/100 g total fatty acid), 10.09 (g/100 g total fatty acid), 35.39, 28.65, and 4.25, respectively. In the wild fish, the EPA level was 6.86, DHA level was 9.28, ∑PUFA level was 32.29, ∑n‐3 level was 24.75, and n‐3/n‐6 ratio was 3.53 [57].

Sea bass (*Dicentrarchus labrax*) is frequently farmed, both in Europe and in Turkey; many studies have focused on this species. Alasavar et al. reported the nutritional compositions of farmed and wild sea bass. In the farmed fish, EPA and DHA values were 6 and 18.1%, respectively, whereas they were 10.06 and 19.5% in the wild fish. In that study, the n‐3/n‐6 ratios were 2.88 and 3.02 in farmed and wild fish, respectively. The researchers attributed the lower levels in farmed fish to SFA and MUFA‐rich, but PUFA‐poor, feeds. They also stated that the wild fish were living in a nutrient‐rich region and fed on n‐3 fatty acid‐rich food sources [58].

Different results were obtained in a study that compared the nutritional compositions of three different fish species (sea bass, sea bream, and rainbow trout) to each other. For sea bass, the EPA level was 7.32%, the DHA level was 14.8%, and the ∑n‐3 level was 26.2%; for sea bream, the levels were 5.48, 12.4, and 23.3%, respectively; for rainbow trout, the levels were 6.16, 19.04, and 31.1%, respectively. The n‐3/n‐6 ratios for each fish species were 3.84, 2.85, and 4.54, respectively. Considering that, in general, the n‐3/n‐6 ratio in a healthy aquaculture food should be at least 1:1, and these three species were well above this value, an indicator of the quality of the aquaculture environment and feeds used in aquaculture [59].

In a study carried out in Turkey, the nutritional compositions of wild fish caught in the Atatürk Dam Lake were investigated: the EPA level was 7.18%, the DHA level was 5.39%, the ∑PUFA level was 23.09%, the ∑n‐3 level was 15.64%, the ∑n‐6 level was 7.45%, and the n‐3/n‐6 ratio was 2.10 [60].

(lean fish; less than 2% fat) from marine water had low lipid content, they were rich in EPA,

There are significant differences in nutritional compositions of farmed fish and wild fish. Many recent studies have focused on this issue and have tried to determine to what degree the nutritional composition of fish affects human health and has nutritional benefits [57–64]. The nutritional quality of farmed fish has improved in the recent years thanks to environmentally friendly and advanced aquaculture techniques. In addition, the advancing feed sector

In its early years, aquaculture was carried out in small areas using artificial feeds and simple techniques, and the only way to handle disease factors was to use antibiotics. However, more recently, aquaculture has improved significantly and has begun to yield quality products that are both environmentally friendly and beneficial to human health. Many studies have shown that the nutritional, and especially, the lipid compositions in farmed fish are more consistent

DHA, and ∑ n‐3 fatty acid relative to the other fish in the same group [56].

**Fish species Fat (g) EPA (g) DHA (g)**

Haddock 1.0 0.07 0.27 Cod 0.6 0.07 0.16 Saithe/coalfish 0.3 – – Tusk 0.3 – – European plaice 1.5 0.24 0.26

Halibut 3.9 0.28 0.41 Atlantic wolffish 2.7 0.40 0.20 Rainbow trout 6.7 0.32 1.16 Spotted wolffish 4.8 0.70 0.40

Greenland Halibut 15.6 1.00 0.90 Salmon 10.0 0.65 1.80 Mackerel 24.4 1.27 3.17 Herring (summer) 14.5 0.57 1.25 Herring (winter) 19.0 2.48 2.24 Eel 31.5 1.27 2.07

than in wild fish; therefore, they are richer in n‐3 fatty acids [59, 63, 65].

now can offer the most suitable and best quality feeds.

**Table 3.** Fat, EPA, and DHA content in selected fish species.

**3.4. Wild or farmed fish**

**Lean fish**

152 Fatty Acids

**Medium fat fish**

**Fatty fish**

Males and females (a total of 10 individual fish) from the shabbout (*Barbus grypus*) species obtained from the same region were used. The samples taken from the species, an omnivorous and rapidly growing species, weighed between 1.5 and 2 kg. Evaluating individual fishes separately showed that EPA values to be between 2.7 and 3.7% and DHA values between 5.2 and 10.7%; the highest value was determined in a male fish. Their ∑PUFA values were between 19.2 and 26.1%; ∑n‐3 values between 14.7 and 18.2%, and, again, the highest value was determined in a male fish. The lowest ∑n‐6 value was 3.9 and determined in a female, whereas the highest ∑n‐6 value was 7.6 and determined in a male fish; thus, at 2.4, the lowest n‐3/n‐6 ratio was determined in male fish and, at 4.8, the highest n‐3/n‐6 was determined in female fish [61].

A similar study was carried out on spiny eel (*Mastacembelus mastacembelus*) and EPA and DHA values were 1.62 and 8.41%, respectively. The ∑PUFA level was 21.74%; ∑n‐3 level, 14.16%, and ∑n‐6 level, 7.11%. The researchers found a n‐3/n‐6 ratio = 2, and asserted that it could be a beneficial species for human health [62]. The researchers asserted that these were the first studies on wild shabbout and spiny eel in the region studied and stated that their results showed that the nutritional and fatty acid compositions of both species were of high quality and can have economic value.

Interesting results were obtained in a study carried out with individual farmed and wild trout. The nutritional compositions of fish obtained from earthen ponds, sea cages, lake (freshwater) cages, and from their natural habitats in lakes were compared. The highest EPA value (8.74%) was found in the wild fish, whereas the lowest EPA value (3.14%) was determined in the lake‐caged fish. To the contrary, at 5.66%, the lowest DHA level was determined in the wild fish and, at 18.49%, the highest DHA value was determined in sea‐caged fish. The ∑n‐3 levels varied between 18.21 and 26.31%; the highest value was determined in sea‐caged fish, the ∑n‐6 values varied between 7.11% (wild fish) and 13.1% (lake‐caged fish). The n‐3/n‐6 ratios were 1.33, 1.75, 2.58, and 2.71 for lake‐caged fish, earthen pond fish, sea‐caged fish, and wild fish, respectively. The n‐3/n‐6 ratios of fish from each different environment were reported to be at acceptable values [63].

### **4. Conclusion**

The nutrients in fish are important for human health, but are easily obtained from fish oils. Fish fatty acids and particularly poly unsaturated fatty acids (PUFA) play an important role in human health, from embryological development to prevention and treatment of some diseases— including arthritis and inflammation, autoimmune disease, type 2 diabetes, hypertension, kidney and skin disorders, and cancer in children and in adults. The human body cannot synthesize certain fatty acids: these essential fatty acids must be consumed in the diet. Therefore, consumption of fish should routinely take place in human nutrition. The fish resources attract consumer interest and have been discussed in detail in the recent years; therefore, many studies have been carried out to investigate the nutritional value of farmed fish. Most of the studies showed that there was no significant difference between farmed and wild fish in terms of nutritional composition. A significant number of these studies mostly focused on the quantity and quality of fish feeds and the edible parts of fish.

### **Author details**

Oğuz Taşbozan\* and Mahmut Ali Gökçe

\*Address all correspondence to: tasbozan@cu.edu.tr

Cukurova University, Fisheries Faculty, Department of Aquaculture, Adana, Turkey

### **References**

[1] Jobling J, Leknes O: Cod liver oil: feed oil influences on fatty acid composition. Aquaculture International. 2010, **18**:223‐230. doi:10.1007/s10499‐008‐9238‐y

[2] Ruxton CHS, Reed SC, Simpson MJA, Millington KJ: The health benefits of omega‐3 polyunsaturated fatty acids: a review of the evidence. Journal of Human Nutrition and Dietetics. 2007, **20**:275‐285. doi:10.1111/j.1365‐277X.2007.00770.x

Interesting results were obtained in a study carried out with individual farmed and wild trout. The nutritional compositions of fish obtained from earthen ponds, sea cages, lake (freshwater) cages, and from their natural habitats in lakes were compared. The highest EPA value (8.74%) was found in the wild fish, whereas the lowest EPA value (3.14%) was determined in the lake‐caged fish. To the contrary, at 5.66%, the lowest DHA level was determined in the wild fish and, at 18.49%, the highest DHA value was determined in sea‐caged fish. The ∑n‐3 levels varied between 18.21 and 26.31%; the highest value was determined in sea‐caged fish, the ∑n‐6 values varied between 7.11% (wild fish) and 13.1% (lake‐caged fish). The n‐3/n‐6 ratios were 1.33, 1.75, 2.58, and 2.71 for lake‐caged fish, earthen pond fish, sea‐caged fish, and wild fish, respectively. The n‐3/n‐6 ratios of fish from each different environment were reported to

The nutrients in fish are important for human health, but are easily obtained from fish oils. Fish fatty acids and particularly poly unsaturated fatty acids (PUFA) play an important role in human health, from embryological development to prevention and treatment of some diseases— including arthritis and inflammation, autoimmune disease, type 2 diabetes, hypertension, kidney and skin disorders, and cancer in children and in adults. The human body cannot synthesize certain fatty acids: these essential fatty acids must be consumed in the diet. Therefore, consumption of fish should routinely take place in human nutrition. The fish resources attract consumer interest and have been discussed in detail in the recent years; therefore, many studies have been carried out to investigate the nutritional value of farmed fish. Most of the studies showed that there was no significant difference between farmed and wild fish in terms of nutritional composition. A significant number of these studies mostly focused on the quantity and quality of fish feeds and the

be at acceptable values [63].

**4. Conclusion**

154 Fatty Acids

edible parts of fish.

**Author details**

**References**

Oğuz Taşbozan\* and Mahmut Ali Gökçe

\*Address all correspondence to: tasbozan@cu.edu.tr

Cukurova University, Fisheries Faculty, Department of Aquaculture, Adana, Turkey

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## **Viable Alternatives Study for Reusing Lipids from Microalgae Biomass Present in the Generated Sludge in the Supply Water Treatment Processes**

Livia de Oliveira Ruiz Moreti, Rosa Maria Ribeiro, Letícia Nishi and Rosângela Bergamasco

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/67484

#### **Abstract**

This chapter aims to evaluate the microalgae species' removal efficiency, using *Moringa oleifera* powder seeds as a natural coagulant with subsequent lipid profile characterization. For the tests were used deionized water artificially contaminated with cell cultures of *Anabaena flos-aquae* and *Chlorella vulgaris*, with a cell density in the order of 10<sup>4</sup> and 10<sup>6</sup> cells mL–1, respectively. Coagulation/flocculation/dissolved air flotation (C/F/DAF) tests were conducted using 'Flotest' equipment, using *M. oleifera* powder seeds in the dosage range of 50–1000 mg L−1. For fatty acid profile analyses, a gas chromatograph equipped with a flame ionization detector was used. Variations of the coagulant dosages showed that there was a difference between dosages and that 100 mg L–1 provided the best removal efficiency for *A. flos-aquae* (96.5, 80.5 and 78.1%) and 140 mg L−1 for *C. vulgaris* (90.5, 78.34 and 70%) of the tested parameters of chlorophyll, color and turbidity, respectively. In relation to the produced sludge, it was observed that the use of this coagulant in the treatment of water contaminated with microalgae produces a biodegradable sludge, rich in lipids, especially oleic acid (>60%). Thus, these results indicate that the sludge's reutilization could be a good alternative to biodiesel production, as it represents an environmentally viable method for reusing residual biomass produced in the water treatment process.

**Keywords:** reusing lipids, microalgae, water treatment processes

### **1. Introduction**

Several natural coagulants have been studied for water purification; recently, *Moringa oleifera* Lam comes to stand out because it has good color and turbidity removal and promotes large

© 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

bacteria removal, of above 90% [1]. Currently, there are few studies on the coagulant application in eutrophic waters, but in some works, can be seen an excellent microalgae cells removal efficiency [2–4].

The dissolved air flotation has been considered a viable alternative to the sedimentation step when applied to water treatment in the microalgae's presence, since this process is capable of removing whole cells, besides decreasing the time contact between the generated sludge with water treatment. The waste generated removal is performed by mechanical equipment, which is installed in water and has an easy maintenance [5, 6].

The proper disposal of the generated sludge by water treatment plants (WTPs) is essential, according to NBR 10004 [7], is considered a solid residue, which if released without proper treatment in waterways, can cause direct effects in the aquatic environment, damaging receiving fauna and flora. The nonchalance towards the waste generated, causes impacts, such as increase in the amount of solids in water body, water body siltation, increased of color, turbidity and aluminum concentration in the water, decrease of the water's pH, releasing of odors, decreases the amount of dissolved oxygen in the water body and chronic toxicity on aquatic organisms and their vision.

However, this chapter seeking environmentally viable alternatives to microalgae biomass reuse present in the generated sludge by water treatment plants, for example, to characterize the lipid content produced by these microalgae, to consign them to a further biodiesel production.

Thus, this chapter shows the removal of *Anabaena flos-aquae* and *Chlorella vulgaris* cells by coagulation/flocculation/dissolved air flotation (C/F/DAF) processes, using *M. oleifera* as a natural coagulant, with subsequent lipid characterization of the generated sludge in order to check the potential for reusing this biomass.

### **2. Microalgae**

Phylogenetically, microalgae are prokaryotes or eukaryotes organisms, according to the period when they appeared on the planet, belonging to a very heterogeneous group of microorganisms. According to Andrade et al. [8] the microalgae are photosynthetic microorganisms that combining water and atmospheric carbon dioxide with sunlight to produce biomass (polysaccharides, proteins, lipids and hydrocarbons). Which can be used in biofuel production, feeding supplements and also can be used in atmospheric carbon dioxide capture. Microalgae produce more oxygen than all plants in the world, accounting for at least 60% of the Earth's primary production.

The microalgae biomass presents about 50% carbon in its composition, so the supply of this nutrient to these microorganisms' cultures represents an important component of the production costs [9]. However, it is necessary to take care of microalgae culture systems, considering the peculiarities of each species, adaptation to the environment, as well as the availability of nutrients associated with economic viability [10].

The exact number of microalgae species is not yet known, but many species may already grow in cropping systems. The most difficult task, however, is to grow specific species for oil production [10].

Some microalgae species under adverse environmental conditions, such as nutrient stress (lack of nitrogen or phosphorus), may accumulate lipids. The green algae specie, *C. vulgaris*, is widely used in research on the biofuel production from microalgae [11, 12], and in the present study, was chosen for the comparison of the lipid content with the cyanobacteria *A. flos-aquae*.

#### **2.1. Microalgae lipid content**

bacteria removal, of above 90% [1]. Currently, there are few studies on the coagulant application in eutrophic waters, but in some works, can be seen an excellent microalgae cells removal

The dissolved air flotation has been considered a viable alternative to the sedimentation step when applied to water treatment in the microalgae's presence, since this process is capable of removing whole cells, besides decreasing the time contact between the generated sludge with water treatment. The waste generated removal is performed by mechanical equipment, which

The proper disposal of the generated sludge by water treatment plants (WTPs) is essential, according to NBR 10004 [7], is considered a solid residue, which if released without proper treatment in waterways, can cause direct effects in the aquatic environment, damaging receiving fauna and flora. The nonchalance towards the waste generated, causes impacts, such as increase in the amount of solids in water body, water body siltation, increased of color, turbidity and aluminum concentration in the water, decrease of the water's pH, releasing of odors, decreases the amount of dissolved oxygen in the water body and chronic toxicity on

However, this chapter seeking environmentally viable alternatives to microalgae biomass reuse present in the generated sludge by water treatment plants, for example, to characterize the lipid content produced by these microalgae, to consign them to a further biodiesel

Thus, this chapter shows the removal of *Anabaena flos-aquae* and *Chlorella vulgaris* cells by coagulation/flocculation/dissolved air flotation (C/F/DAF) processes, using *M. oleifera* as a natural coagulant, with subsequent lipid characterization of the generated sludge in order to

Phylogenetically, microalgae are prokaryotes or eukaryotes organisms, according to the period when they appeared on the planet, belonging to a very heterogeneous group of microorganisms. According to Andrade et al. [8] the microalgae are photosynthetic microorganisms that combining water and atmospheric carbon dioxide with sunlight to produce biomass (polysaccharides, proteins, lipids and hydrocarbons). Which can be used in biofuel production, feeding supplements and also can be used in atmospheric carbon dioxide capture. Microalgae produce more oxygen than all plants in the world, accounting for at least

The microalgae biomass presents about 50% carbon in its composition, so the supply of this nutrient to these microorganisms' cultures represents an important component of the production costs [9]. However, it is necessary to take care of microalgae culture systems, considering the peculiarities of each species, adaptation to the environment, as well as the

is installed in water and has an easy maintenance [5, 6].

aquatic organisms and their vision.

check the potential for reusing this biomass.

60% of the Earth's primary production.

availability of nutrients associated with economic viability [10].

efficiency [2–4].

162 Fatty Acids

production.

**2. Microalgae**

The microalgae biomass contains three main components: carbohydrates, proteins and lipids [13]. In biological systems, the lipids function as membrane components, reserve products, metabolites and as energy sources, with most of them consists of fatty acids. Thus, the lipids are classified in storage lipids (neutral lipids), triacylglycerols and membrane lipids (polar lipids), phospholipids, glycolipids and sterols [14].

Fatty acids are fundamental units most of the lipids. They are short-chain and long-chain organic acids having 4–24 carbon atoms, and short-chain fatty acids are ideal for the biodiesel production [15].

Some fatty acids synthesized by microalgae, such as omega 3 and 6 (ω-3 and ω-6), which are the main precursors of some hormones such as prostaglandins, prostacyclins, leukotrienes and thromboxanes, have a high economic value in the food and pharmaceutical industry and are fundamental for the development and physiological regulators [16].

Fatty acids in microalgae correspond to the largest lipid fraction, and, in some species, polyunsaturates represent between 25 and 60% of total lipids [17].

The polyunsaturated fatty acids from microalgae have a very promising market in biotechnology, especially in the functional food industry [18]. Studies presented by Favaro-Trindade et al.(2008) [48] show that lipids, especially unsaturated fatty acids, have been encapsulated to reduce susceptibility to oxidation.

According to Nelson e Cox [14], fatty acids have a unique carboxyl group and a non-polar hydrocarbon tail, which give lipids their oily and fatty nature, insoluble in water. They occur in cells or tissues in forms covalently attached to different lipids' classes. Different fatty acids have been isolated from lipids of various species.

They differ by the chain extension and its presence, number and double bonds position, and some fatty acids also have methyl-branched groups.

The glycolipids that are composed of glycerol have been found in many organisms, being observed as the main lipid component of microalgae photosynthetic membranes, including cyanobacteria (blue microalgae). Its structure is analogous to that of glycerophospholipids with a sugar molecule, glycosidically attached to glycerol three position and fatty acids esterified in the other two positions.

Among the main glycosylacylglycerols of microalgae and plant photosynthetic membranes is monogalactosyl-diacylglycerol (MGDG), which occurs abundantly in plants and algae, especially in chloroplasts. Contains high proportions of polyunsaturated fatty acids. For *C. vulgaris*, the MGDG presents mainly oleic acid (C18:1) and linoleic acid (C18:2) when cultivated in the dark and 20% of linolenic acid (C18:3) when cultivated in the light [19].

The lipid production estimation by microalgae ranges from 15,000 to 30,000 L.km<sup>2</sup> , and its extraction is simple and can be applied to traditional methods used in the chemical industry, including solvent extraction [20].

It is known that among the nutrients that can influence the lipids and fatty acids production are the sources of nitrogen and sulfur, which are used by microalgae in the synthesis of amino acids and fatty acids [20].

The main applications of fatty acids microalgae occur in the enrichment of fish feed, biodiesel production and sources of essential unsaturated fatty acids in the human diet [21].

Although there are many microorganism types capable to accumulating lipids, not all of them have favorable characteristics for the application in the biodiesel production. The microalgae stand out because they present, in some cases, compatibility in the ratio of their oil produced to the vegetable oil used in the transesterification process [22, 23].

According to Schimitz et al. [10], the presence of polyunsaturated compounds produced by microalgae causes a decrease in the stability of produced biodiesel. However, due to the presence of these fatty acids, biodiesel from microalgae presents a high yield at low temperatures, a characteristic that is not presented by conventional oilseed biodiesel, which have little yield at relatively low temperatures.

#### **2.2.** *Moringa oleifera* **Lam**

*M. oleifera* Lam is a tropical tree that grows naturally in India, South Saharan Africa and South America [24]. The leaves, flowers, seeds, roots and bark may be used as food or for medicinal and therapeutic purposes [25], especially in developing countries [24, 26]. In addition, other applications were pointed out for cosmetics preparation, mechanical lubricants and even for potential biodiesel elaboration [27].

According to Ndabigengesere and Narasiah e Talbot [28], *M. oleifera* seeds contain about 37% protein, 35% lipids and 5% carbohydrates (oligosaccharides). The carbohydrate content is very low whereas the high lipid content explains why the seeds can be used as a source of vegetable oil. This oil resembles olive oil in its composition, being rich in oleic acid, which makes it suitable for edible purposes [29].

The *M. oleifera* seeds are also very useful as a coagulant in drinking water clarification and effluent treatment since 1979, due to the presence of a water soluble cationic coagulant protein capable of reducing the turbidity of the treated water. The seeds can be used in the form of powder, such as aqueous or saline extracts [24, 30, 31].

#### **2.3. Removal of microalgae from water using** *Moringa oleifera***, as natural coagulant**

Among the main glycosylacylglycerols of microalgae and plant photosynthetic membranes is monogalactosyl-diacylglycerol (MGDG), which occurs abundantly in plants and algae, especially in chloroplasts. Contains high proportions of polyunsaturated fatty acids. For *C. vulgaris*, the MGDG presents mainly oleic acid (C18:1) and linoleic acid (C18:2) when cultivated in the dark and 20% of linolenic acid (C18:3) when cultivated in

extraction is simple and can be applied to traditional methods used in the chemical industry,

It is known that among the nutrients that can influence the lipids and fatty acids production are the sources of nitrogen and sulfur, which are used by microalgae in the synthesis of amino

The main applications of fatty acids microalgae occur in the enrichment of fish feed, biodiesel

Although there are many microorganism types capable to accumulating lipids, not all of them have favorable characteristics for the application in the biodiesel production. The microalgae stand out because they present, in some cases, compatibility in the ratio of their oil produced

According to Schimitz et al. [10], the presence of polyunsaturated compounds produced by microalgae causes a decrease in the stability of produced biodiesel. However, due to the presence of these fatty acids, biodiesel from microalgae presents a high yield at low temperatures, a characteristic that is not presented by conventional oilseed biodiesel, which have little yield

*M. oleifera* Lam is a tropical tree that grows naturally in India, South Saharan Africa and South America [24]. The leaves, flowers, seeds, roots and bark may be used as food or for medicinal and therapeutic purposes [25], especially in developing countries [24, 26]. In addition, other applications were pointed out for cosmetics preparation, mechanical lubricants and even for

According to Ndabigengesere and Narasiah e Talbot [28], *M. oleifera* seeds contain about 37% protein, 35% lipids and 5% carbohydrates (oligosaccharides). The carbohydrate content is very low whereas the high lipid content explains why the seeds can be used as a source of vegetable oil. This oil resembles olive oil in its composition, being rich in oleic acid, which

The *M. oleifera* seeds are also very useful as a coagulant in drinking water clarification and effluent treatment since 1979, due to the presence of a water soluble cationic coagulant protein capable of reducing the turbidity of the treated water. The seeds can be used in the form of

, and its

The lipid production estimation by microalgae ranges from 15,000 to 30,000 L.km<sup>2</sup>

production and sources of essential unsaturated fatty acids in the human diet [21].

to the vegetable oil used in the transesterification process [22, 23].

the light [19].

164 Fatty Acids

including solvent extraction [20].

acids and fatty acids [20].

at relatively low temperatures.

potential biodiesel elaboration [27].

makes it suitable for edible purposes [29].

powder, such as aqueous or saline extracts [24, 30, 31].

**2.2.** *Moringa oleifera* **Lam**

Coagulation/flocculation (C/F) followed by dissolved air flotation (DAF) is suitable for the treatment of naturals and synthetic eutrophic waters [5, 32]. When it comes to the removal of cyanobacterial cells, DAF is an effective alternative, as shown by some studies in the literature [5, 6]. However, to achieve good efficiency, water treatment plants (WTPs) use a series of auxiliary products in the process, especially the use of inorganic coagulants, usually based on metals such as aluminum, as well as pH control. However, these coagulants do not generate biodegradable sludge, causing problems in terms of disposal and treatment; this may be also related to some diseases, such as Alzheimer's disease, due to residual aluminum in treated water [33, 34]. Thus, the search becomes necessary for alternative natural coagulants that are biodegradable and safe to human health [35].

The *M. oleifera* (MO) seeds can be used for efficient clarification of water [26, 36]. This efficiency has been shown by high values of color, turbidity and bacteria removal [2, 37] and even cyanobacteria cells in the water treatment process [38], as well as some economic and environmental advantages related to decreasing the costs of synthetic products to correct the pH of water and produce a sludge without metals.

The water treatment processes in WTPs produce residues, mostly water used for washing the filters and sludge from sedimentation tanks/floaters [39]. Particularly, in WTPs with cyanobacteria problems, the sludge generated is composed of microalgae biomass. Knowing that such biomass has a relatively high amount of lipids in their composition [40], which could be used for biodiesel production.

Firstly, by evaluating the results obtained (**Figure 1**), one can observe the percentages of the removal of the parameters color, turbidity and chlorophyll-a and compounds with absorp-

**Figure 1.** Color, turbidity, compounds with absorption in UV254 nm and chlorophyll-a removal in relation to the dosage of *Moringa oleifera* powder seeds for *Anabaena flos-aquae*.

tion in UV254 nm, indicating the optimum dosage of the *M. oleifera* powder seeds used in water contaminated with cyanobacteria species (*A. flos-aquae*).

The results indicate that the *M. oleifera* powder seed added directly to cell suspensions was effective in removing cells, color and turbidity, reaching up to 96.4, 80.5 and 78.1%, respectively, for the dosage of 100 mg L−1, which was considered ideal for this study.

Regarding the removal of UV254 nm, it was observed that *M. oleifera* didn't obtain very satisfactory results, reaching 39.1% removal in the dosage of 100 mg L−1. There was a drop in the removal efficiency as the *M. oleifera* dosage was increased. This result can be attributed to the fact that *M. oleifera* is an organic coagulant, basically composed of proteins, lipids and carbohydrates, responsible for the organic residual in the treated water.

The optimum *M. oleifera* dosage for the *C. vulgaris* species, which is a unicellular microalga, was also evaluated in order to verify if the different morphology of the microalgae interferes in the parameters of removal efficiency. In this way, it can be observed that the optimum coagulant dosage was different among the species.

For *C. vulgaris*, the optimum coagulant dosage was 400 mg L−1, verified by the removal efficiency for color (78.34%), turbidity (70%), chlorophyll-a (90.5%) and UV254 nm (16%) absorption compounds as shown in **Figure 2**.

Thus, it was observed that the C/F/DAF processes used together with *M. oleifera* as a coagulant had an excellent efficiency for both species that were tested.

In relation to the microalgae lipid profile analysis, the fatty acids and esters microalgae were first identified without the *M. oleifera* treatment.

**Figure 2.** Color, turbidity, compounds with absorption in UV254 nm and chlorophyll-a removal in relation to the dosage of *Moringa oleifera* powder seeds for *Chlorella vulgaris*.

It can be verified that the saturated fatty acids corresponded to 40.4% composition of *C. vulgaris* and 35.85% of *A. flos-aquae*, whereas the unsaturated ones presented values of 39.58 and 40.1%, respectively.

tion in UV254 nm, indicating the optimum dosage of the *M. oleifera* powder seeds used in water

The results indicate that the *M. oleifera* powder seed added directly to cell suspensions was effective in removing cells, color and turbidity, reaching up to 96.4, 80.5 and 78.1%, respec-

Regarding the removal of UV254 nm, it was observed that *M. oleifera* didn't obtain very satisfactory results, reaching 39.1% removal in the dosage of 100 mg L−1. There was a drop in the removal efficiency as the *M. oleifera* dosage was increased. This result can be attributed to the fact that *M. oleifera* is an organic coagulant, basically composed of proteins, lipids and carbo-

The optimum *M. oleifera* dosage for the *C. vulgaris* species, which is a unicellular microalga, was also evaluated in order to verify if the different morphology of the microalgae interferes in the parameters of removal efficiency. In this way, it can be observed that the optimum

For *C. vulgaris*, the optimum coagulant dosage was 400 mg L−1, verified by the removal efficiency for color (78.34%), turbidity (70%), chlorophyll-a (90.5%) and UV254 nm (16%) absorption

Thus, it was observed that the C/F/DAF processes used together with *M. oleifera* as a coagulant

In relation to the microalgae lipid profile analysis, the fatty acids and esters microalgae were

color Turbidity Chlorophyll-a

UV254

50 100 200 400 500 600 800 1000

*Moringa oleifera dosages (mg.L-1)* 

**Figure 2.** Color, turbidity, compounds with absorption in UV254 nm and chlorophyll-a removal in relation to the dosage of

tively, for the dosage of 100 mg L−1, which was considered ideal for this study.

hydrates, responsible for the organic residual in the treated water.

coagulant dosage was different among the species.

first identified without the *M. oleifera* treatment.

*Moringa oleifera* powder seeds for *Chlorella vulgaris*.

had an excellent efficiency for both species that were tested.

compounds as shown in **Figure 2**.


0

20

40

**Removal (%)** 

60

80

100

166 Fatty Acids

contaminated with cyanobacteria species (*A. flos-aquae*).

Among the acids with the highest values in *C. Vulgaris* were C20:0 (arachidic acid) with 21.15%, C18:1n9 (oleic acid) with 18.85% followed by C16:0 (palmitic acid) and C18:2n6 (linoleic acid) with about 15% each. Already *A. flos-aquae*, the highest percentages are due first to C16:0 (palmitic acid) with 30.55%, then C18:2n6 (linoleic acid) presented 17% and, finally, C18: 1n9 (oleic acid) presented 7.4% of its composition, shown in **Table 1**.



**Table 1.** Microalgae chromatographic profile without *Moringa oleifera* treatment.

The species *C. vulgaris* presented a lipid content of 5% of its dry weight, a value higher than the cyanobacteria studied (*A. flos-aquae*), which was 3.05%, but this result can be reversed, since the medium and conditions as potentiated as lipid productions by microalgae, as light [41], carbon dioxide concentration, temperature [42, 43], nitrogen source concentration [43], among other nutrients. The value obtained for *C. vulgaris* is in agreement with the results obtained by Radman and Costa [20], which presented concentrations of approximately 5.97% of lipid content for the same microalgae under the same culture conditions.

Embora haja poucos trabalhos relatando o perfil lipídico de *Anabaena flos-aquae*, os resultados obtidos neste estudo apresentam valores próximos aos apresentados por Nichols e Wood (1967) em que apresentam C16:0 (39,5%), C16:1 (5,5%), C18:1 (5,2%) C18:2 (36,5%) como os principais ácidos graxos pertencentes da maioria da composição desta alga.

Although there are few studies reporting the *A. flos-aquae* lipid profile, the results obtained in this chapter present values close to those presented by Nichols and Wood [44] in which they present C16:0 (39.5%), C16:1 (5.5%), C18:1 (5.2%) and C18:2 (36.5%) as the main fatty acids belonging to the majority of this algae composition.

After treatment with the *M. oleifera* optimal dosages, the analysis was repeated, and it was observed that results presented in **Table 2** demonstrate that after the treatment with the coagulant optimal dosages, the total lipid percentages of each sample suffered an increase, seen by the values of 16.4% of the total lipids for *C. vulgaris* and 6.2% for *A. flos-aquae*.

This increase is probably related to the residual *M. oleifera* coagulant in the sludge because, according to the *M. oleifera* seeds' physicochemical characterization from Aracaju–SE, they present approximately 37% of lipids in their composition. Therefore, most of the fatty acids present in the samples for both microalgae species were unsaturated fatty acids.

The acid responsible for this increase was C18:1n9 (oleic acid), presenting 69.5% in the *C. vulgaris* sample against 61.7% in the *A. flos-aquae* sample. These results are in agreement with those reported by Silva et al. [45] in which the oil extracted from *M. oleifera* is characterized and with the presence of 78% of oleic acid. Rashid et al. [46] also presented more than 70% of oleic acid in its *Moringa* samples. According to him, some oscillations in the fatty acids' values can occur related to the conditional variations used by the farmers such as fertilizers, soil and the seed variety.

Viable Alternatives Study for Reusing Lipids from Microalgae Biomass Present in the Generated Sludge in the Supply... http://dx.doi.org/10.5772/67484 169


**Table 2.** Microalgae chromatographic profile after treatment with *Moringa oleifera*.

The species *C. vulgaris* presented a lipid content of 5% of its dry weight, a value higher than the cyanobacteria studied (*A. flos-aquae*), which was 3.05%, but this result can be reversed, since the medium and conditions as potentiated as lipid productions by microalgae, as light [41], carbon dioxide concentration, temperature [42, 43], nitrogen source concentration [43], among other nutrients. The value obtained for *C. vulgaris* is in agreement with the results obtained by Radman and Costa [20], which presented concentrations of approximately 5.97%

**treatment**

**Total** 100 100

**Fatty acids** *Chlorella vulgaris Anabaena flos-aquae*

Subtotal 37.23 36.2 Not identified 19.8 23.95

Total lipids 5% 3.05%

**Table 1.** Microalgae chromatographic profile without *Moringa oleifera* treatment.

168 Fatty Acids

**% Fatty acids means present in microalgae without** 

Embora haja poucos trabalhos relatando o perfil lipídico de *Anabaena flos-aquae*, os resultados obtidos neste estudo apresentam valores próximos aos apresentados por Nichols e Wood (1967) em que apresentam C16:0 (39,5%), C16:1 (5,5%), C18:1 (5,2%) C18:2 (36,5%) como os

Although there are few studies reporting the *A. flos-aquae* lipid profile, the results obtained in this chapter present values close to those presented by Nichols and Wood [44] in which they present C16:0 (39.5%), C16:1 (5.5%), C18:1 (5.2%) and C18:2 (36.5%) as the main fatty acids

After treatment with the *M. oleifera* optimal dosages, the analysis was repeated, and it was observed that results presented in **Table 2** demonstrate that after the treatment with the coagulant optimal dosages, the total lipid percentages of each sample suffered an increase, seen by

This increase is probably related to the residual *M. oleifera* coagulant in the sludge because, according to the *M. oleifera* seeds' physicochemical characterization from Aracaju–SE, they present approximately 37% of lipids in their composition. Therefore, most of the fatty acids

The acid responsible for this increase was C18:1n9 (oleic acid), presenting 69.5% in the *C. vulgaris* sample against 61.7% in the *A. flos-aquae* sample. These results are in agreement with those reported by Silva et al. [45] in which the oil extracted from *M. oleifera* is characterized and with the presence of 78% of oleic acid. Rashid et al. [46] also presented more than 70% of oleic acid in its *Moringa* samples. According to him, some oscillations in the fatty acids' values can occur related to the conditional variations used by the farmers such as fertilizers, soil and the seed variety.

of lipid content for the same microalgae under the same culture conditions.

principais ácidos graxos pertencentes da maioria da composição desta alga.

the values of 16.4% of the total lipids for *C. vulgaris* and 6.2% for *A. flos-aquae*.

present in the samples for both microalgae species were unsaturated fatty acids.

belonging to the majority of this algae composition.

According to Qu et al. [47], oils with high oleic acid values (>70%) improve the biodiesel properties, such as cold flow, cloud point and pour point, in this way, the sludge produced after treatment with the *M. oleifera* optimum dosage presented high oleic acid values in its composition, data that make interesting the use of this residue to a future production of biodiesel.

### **Author details**

Livia de Oliveira Ruiz Moreti1 , Rosa Maria Ribeiro<sup>2</sup> , Letícia Nishi1 \* and Rosângela Bergamasco1


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treatment with the *M. oleifera* optimum dosage presented high oleic acid values in its composition, data that make interesting the use of this residue to a future production of biodiesel.

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\*Address all correspondence to: leticianishi@hotmail.com

1 State University of Maringa, Maringá-PR, Brasil

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**Fatty Acids in Veterinary and Dairy Products**

## **Fatty Acids in Veterinary Medicine and Research**

Siobhan Simpson, Alison Mostyn and

Catrin S. Rutland

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/intechopen.68440

#### **Abstract**

Fatty acid regulation is an essential process for all animals. A number of studies have shown that diet affects the levels/availability of fatty acids in the body but increasingly an evidence shows that disease states can alter the amounts within the body too. Fatty acid levels and availability have been altered by a number of diseases, disorders and reactions including inflammatory responses, heart disease and heart failure and wound repair. They are also essential during the growth and development stages of animals. The amount of research into the consequences of different fatty acid intake and levels in various disease states and during development has increased in both humans and animals. This review presents an overview of the research undertaken to date and highlights the importance, uses and benefits of understanding the roles of fatty acids in both the healthy animals and animals under differing disorders and diseases.

**Keywords:** heart disease, Inflammation, development, nutrition, cancer, pregnancy

### **1. Introduction to fatty acids**

Fatty acids consist of a carboxylic acid with a hydrocarbon chain tail, the length of which varies between fatty acids, as does the presence or absence of double bonds between the carbon atoms and their location [1]. Some fatty acids are ingested in the diet whereas others are synthesized into other fatty acids by elongation and desaturation enzymes [2–4], see **Figures 1** and **2**. In mammals, fatty acids are obtained from the diet prior to metabolism or incorporation as components of cells [5–8]. *n*-6 polyunsaturated fatty acids (PUFAs) and *n*-3 PUFAs are the two major groups of fatty acids; the first is obtained from fats and oils, and the latter from

© 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

**Figure 1.** Schematic of linoleic and arachidonic acid biosynthetic pathway derived from KEGG pathway maps [2].

fish and seafood products [6]. It is essential that the precursors of both *n*-6 and *n*-3 PUFAs are extracted by mammals from their diet as they are not able to convert these fatty acids (FAs) between the two major pathways [9].

**Figure 2.** Schematic of palmitic and oleic acid biosynthesis pathway derived from KEGG pathway maps [2].

### **2. Inflammation, disease and the immune system**

fish and seafood products [6]. It is essential that the precursors of both *n*-6 and *n*-3 PUFAs are extracted by mammals from their diet as they are not able to convert these fatty acids (FAs)

**Figure 1.** Schematic of linoleic and arachidonic acid biosynthetic pathway derived from KEGG pathway maps [2].

between the two major pathways [9].

178 Fatty Acids

Fatty acids are crucial components of the immune system, providing the structural basis of all cell membranes, acting as signaling molecules, and providing a major substrate for energy production [1, 8, 10]. Many diseases involve inflammatory responses either as a reaction to disease or in the initiation of the disease process; although inflammation itself is not always detrimental, for instance, it is an important aspect of wound repair [11–14]. Elevated markers of inflammation are frequently detected in heart failure and cancers although this could be due to the response to disease, or the underlying cause of disease [15–19].

Fatty acid-derived eicosanoids are important contributors to the inflammatory response [13, 20, 21]. The *n*-6 PUFA arachidonic acid is a precursor of the most important pro-inflammatory eicosanoids, while the *n*-3 PUFA derivatives, eicosapentaenoic acid and docosahexaenoic acid metabolites are considered less inflammatory [20]. Arachidonic acid is released from cell membranes by phospholipase A<sup>2</sup> enzymes in response to pro-inflammatory stimuli [22–25]. Cyclooxygenase, lipoxygenase and cytochrome P450 enzymes then convert free arachidonic acid into eicosanoids [26–29]; however, these enzymes are rate limiting as they similarly convert other fatty acids to their metabolites [20]. It has been suggested that if cyclooxygenase, lipoxygenase and cytochrome P450 enzymes are exposed to increased levels of *n*-3 fatty acids, the result is fewer arachidonic acid-derived eicosanoids [20, 30].

Due to the difference in the inflammatory response between fatty acid metabolites, it is hypothesized that the fatty acid profiles could differ between diseased and healthy individuals. Indeed, fatty acid profiles have been shown to be altered in blood and tissues in individuals with a range of conditions compared to unaffected individuals in both humans and dogs. These conditions include Crohn's disease, heart disease, skin disease and cancer [31–34], and are discussed in greater detail below.

#### **2.1. The role of fatty acids in Crohns' disease**

An interesting inflammatory response disorder is inflammatory bowel disease, including Crohn's disease. A number of animal studies, including guinea pigs and rats, have shown novel results in the adipocytes, lipid rafts and fatty acid-derived messenger molecules which indicated that aberrant fatty acid composition could play a role in Crohn's disease [35–38]. This research led directly into looking at the role of FAs in human cases of Crohns' disease, a disorder which is linked to both inflammation and the immune system. Perinodal adipose tissue (PAT) is a specialized adipose tissue depot which surrounds lymph nodes and acts in a paracrine manner—delivering specific FAs and adipokines directly to the node. Research has demonstrated that PAT associated with the lymph node is present in most animals and humans [39]. Crohn's disease is associated with altered mesenteric PAT FA content, suggesting impaired delivery of FAs to lymphocytes [40]. For many years, patients with Crohn's disease have been advised to take dietary fish oils that are rich in *n*-3 PUFAs, but interestingly patients have naturally (prior to taking supplements) presented with higher levels of *n*-3 PUFAs than observed in controls with concurrent deficiencies in arachidonic acid (20:4*n-*6) [41–43]. More recent evidence suggests that higher levels of *n*-6 PUFAs, including linoleic acid (18:2*n*-6) were most effective at relieving inflammatory symptoms [43]. The biosynthetic links between arachidonic acid (20:4*n-*6) and linoleic acid (18:2*n*-6) are shown in **Figure 1** and help to understand why an increased linoleic acid intake could reverse the decrease in arachidonic acids observed in patients. A number of animal species develop differing forms of inflammatory bowel disease, therefore understanding whether FAs are affected for the differing types of animals and differing breeds could help to indicate differing dietary or treatment requirements.

#### **2.2. The role of fatty acids in cardiovascular function and disease**

A number of links have been made between fatty acid levels and heart disease and heart failure. Human patients with significant left ventricular dilation have a larger percentage of oleic acid and a smaller percentage of arachidonic acid in their blood serum compared to patients with moderate left ventricular dilation [33]. It is also important to highlight that none of the patients involved in the study had a confirmed diagnosis, and although valve disease and coronary artery disease were excluded as the underlying cause of left ventricular dilation, infarction was not. Infarction may have skewed the fatty acid results due to the strong inflammatory nature of myocardial infarction [33, 44].

In cats with hypertrophic cardiomyopathy, differing levels of FAs were observed when compared to cats with no hypertrophic symptoms [45]. Hypertrophic cardiomyopathy cats had higher levels of docosahexaenoic acid, palmitic acid and total *n*-3 PUFAs and lower levels of linoleic acid. Differential levels of docosatetraenoic acid have been observed in canine myocardial tissue in dogs affected by dilated cardiomyopathy [46]. Mobile lipid content within the myocardium was significantly increased in a 24-hour coronary occlusion canine heart, not only throughout the body but also 'local' increases were observed around the heart with cardiac levels up to 10 times higher than the rest of the body [47–50]. It has been suggested that increased fatty acid levels alongside a decrease in creatine can lead to diastolic dysfunction, as observed in humans with diabetic cardiomyopathy [51, 52]. Despite the observations in dogs and humans, a study in rats showed increased fatty acids and decreased creatine but no associated diastolic dysfunction was observed [53]. With differing observations between species, more research is needed in order to understand the mechanisms and circumstances under which diastolic alteration occurs. Increased levels of palmitoleic acid have been associated with heart failure, higher levels of behenic acid and stearic acid have been associated with lower risk of developing atrial fibrillation, women with higher circulating pentadecanoic acid are less likely to have a myocardial infarction, hypertensive rats have higher circulating eicosedienoic acid and in renal patients higher circulating C20:5n3 is associated with good cardiac functional measures [54–60].

similarly convert other fatty acids to their metabolites [20]. It has been suggested that if cyclooxygenase, lipoxygenase and cytochrome P450 enzymes are exposed to increased levels

Due to the difference in the inflammatory response between fatty acid metabolites, it is hypothesized that the fatty acid profiles could differ between diseased and healthy individuals. Indeed, fatty acid profiles have been shown to be altered in blood and tissues in individuals with a range of conditions compared to unaffected individuals in both humans and dogs. These conditions include Crohn's disease, heart disease, skin disease and cancer [31–34], and

An interesting inflammatory response disorder is inflammatory bowel disease, including Crohn's disease. A number of animal studies, including guinea pigs and rats, have shown novel results in the adipocytes, lipid rafts and fatty acid-derived messenger molecules which indicated that aberrant fatty acid composition could play a role in Crohn's disease [35–38]. This research led directly into looking at the role of FAs in human cases of Crohns' disease, a disorder which is linked to both inflammation and the immune system. Perinodal adipose tissue (PAT) is a specialized adipose tissue depot which surrounds lymph nodes and acts in a paracrine manner—delivering specific FAs and adipokines directly to the node. Research has demonstrated that PAT associated with the lymph node is present in most animals and humans [39]. Crohn's disease is associated with altered mesenteric PAT FA content, suggesting impaired delivery of FAs to lymphocytes [40]. For many years, patients with Crohn's disease have been advised to take dietary fish oils that are rich in *n*-3 PUFAs, but interestingly patients have naturally (prior to taking supplements) presented with higher levels of *n*-3 PUFAs than observed in controls with concurrent deficiencies in arachidonic acid (20:4*n-*6) [41–43]. More recent evidence suggests that higher levels of *n*-6 PUFAs, including linoleic acid (18:2*n*-6) were most effective at relieving inflammatory symptoms [43]. The biosynthetic links between arachidonic acid (20:4*n-*6) and linoleic acid (18:2*n*-6) are shown in **Figure 1** and help to understand why an increased linoleic acid intake could reverse the decrease in arachidonic acids observed in patients. A number of animal species develop differing forms of inflammatory bowel disease, therefore understanding whether FAs are affected for the differing types of animals and differing breeds could help to indicate differing dietary or treatment requirements.

of *n*-3 fatty acids, the result is fewer arachidonic acid-derived eicosanoids [20, 30].

are discussed in greater detail below.

180 Fatty Acids

**2.1. The role of fatty acids in Crohns' disease**

**2.2. The role of fatty acids in cardiovascular function and disease**

matory nature of myocardial infarction [33, 44].

A number of links have been made between fatty acid levels and heart disease and heart failure. Human patients with significant left ventricular dilation have a larger percentage of oleic acid and a smaller percentage of arachidonic acid in their blood serum compared to patients with moderate left ventricular dilation [33]. It is also important to highlight that none of the patients involved in the study had a confirmed diagnosis, and although valve disease and coronary artery disease were excluded as the underlying cause of left ventricular dilation, infarction was not. Infarction may have skewed the fatty acid results due to the strong inflamAlthough the fatty acids themselves play a key role in cardiovascular health and disease, other molecules within the fatty acid utilization cascades play important roles too. Heart-type fatty acid-binding protein (H-FABP) is expressed in cardiomyocytes and despite the name, it is also expressed in renal and skeletal muscle cells [61]. Heart-type fatty acid-binding protein (H-FABP) is used as a prognosis tool biomarker in human cardiac disease as it indicates myocardial stretch and injury in chronic heart failure even in children. Higher levels of H-FABP are associated with a poorer long-term outcome in both adults and children [61–65]. Although little work has been carried out in other species, this is an area of research which has potential, in addition to investigating whether H-FABP levels are raised prior to infarction and/or heart disease. A rat model has shown that H-FABP is increased following cardiac injury [66]. It also enables detection via a number of differing methods including EIA, ELISA, fully automated latex-agglutination assay and qualitative lateral-flow assay microparticle enhanced immunoassay [61].

External factors such as diet and surgery can play large roles in fatty acid composition and cardiovascular health. A study looking at differing feeding regimes in obese rats in comparison with lean rats showed that *n*-3 acyl chains, unsaturated and polyunsaturated fatty acids, were all significantly higher in *obese* rats than in the *lean* ones [53]. What was also interesting was the fact that mild, short-term diet changes (food intake was restricted by 20% for two weeks) did not alter the cardiac fatty acid profiles. The obese mice also showed symptoms of early stage obese cardiomyopathy; although interestingly the symptoms of this started to improve upon calorie restriction, an important finding as it showed that mild calorie restriction can be of benefit under these circumstances. Fatty acids are not only an important indicator of heart disease in animals, but also important in situations such as surgery. Increased free fatty acid levels also have been noted in response to heart surgery in pigs especially when heparin is co-administered [67]. In the surgery cases, it was found that the young patients were more affected than older patients and the levels were more likely to rise if cyanosis and prolonged ischemia were present.

Although most of the work into cardiovascular health has concentrated on disease and disorders, a number of suggestions for healthy levels have been put forward as ways of preventing disease. There is some evidence that higher levels of circulating arachidic acid are associated with lower risk of atrial fibrillation and diabetes [57, 68]. Another example is docosahexaenoic acid (*n*-3 PUFA) which has been implicated as having beneficial effects in a wide range of diseases including heart disease and neurological dysfunction [55, 69].

#### **2.3. Fatty acids and skin disease**

There are two main ways in which differing fatty acid profiles contribute to skin disease—as part of inflammation and affecting membrane fluidity. These are not mutually exclusive and it is possible that fatty acids are affecting the development of skin disease via both. People with atopic eczema have been shown to have a different fatty acid profile in their skin than people without atopic eczema. In particular, they have shorter fatty acids within their skin than unaffected individuals. This difference is suggested to lead to impaired skin barrier [70]. Atopic eczema is an inflammatory disease and thus processes of inflammation as discussed earlier will be active in the disease process [71]. As with other cases where a difference in fatty acid profiles has been established between individuals with disease and healthy individuals, it is not clear whether the fatty acid change causes the disease or is a response to disease, or possibly both, but it is a potential novel treatment route. Similar to people with atopic eczema, pruritic dogs have been shown to have a different fatty acid profile compared to dogs with healthy skin [72]. More recently, dogs with atopic dermatitis whose diets were supplemented with *n*-3 PUFA improved significantly more than those given the placebo [73]. As with human skin disease, it is not clear as to how this works, but it is an additional treatment option and area for further research.

#### **2.4. Cancer associations with fatty acids**

Cancer is the result of aberrant cellular processes. Many genes and proteins are differentially expressed in tumor tissue compared to nontumor tissue [74–77]. Thus, it is intuitive that fatty acid profiles are likely to be altered in tumors compared to nontumor tissue and this has indeed been demonstrated in breast and prostate cancer [78, 79].

There have been studies showing that differential dietary intake of fatty acids can either reduce or increase risk of disease, including cancer. A meta-analysis of studies relating breast cancer risk with *n*-3 PUFA intake showed that overall increasing *n*-3 PUFA intake reduced the risk of developing breast cancer [78]. In transgenic mice in which males develop prostate cancer, *n*-3 PUFA intake from marine sources suppressed tumorigenesis [80]. This is also the case in people where there is reduced risk of developing prostate cancer with increased intake of marine *n*-3 PUFAs [81–83]. Longer chain *n*-3 PUFAs from non-marine sources, however, are associated with an increased risk of prostate cancer [79, 82, 83].

While ultimately work is required in whole organisms, cell lines are a valuable starting point for research. Of particular note in relation to veterinary medicine and fatty acids are two studies on canine tumor cell lines. The first is that of canine lymphoma cell lines; in this study, stearidonic acid was shown to sensitize cells to anticancer drugs, even when the cells were previously resistant to drugs [84]. The second study utilized fatty acids themselves as antitumor agents. In this study, a specific fatty acid, *trans*-10, *cis*-12 conjugated linoleic acid, was shown to inhibit cell growth and induce apoptosis in canine osteosarcoma cell lines and canine lipomas [85, 86].

### **3. The effects of fatty acids on fertility and during pregnancy and development**

Although most of the work into cardiovascular health has concentrated on disease and disorders, a number of suggestions for healthy levels have been put forward as ways of preventing disease. There is some evidence that higher levels of circulating arachidic acid are associated with lower risk of atrial fibrillation and diabetes [57, 68]. Another example is docosahexaenoic acid (*n*-3 PUFA) which has been implicated as having beneficial effects in a wide range of dis-

There are two main ways in which differing fatty acid profiles contribute to skin disease—as part of inflammation and affecting membrane fluidity. These are not mutually exclusive and it is possible that fatty acids are affecting the development of skin disease via both. People with atopic eczema have been shown to have a different fatty acid profile in their skin than people without atopic eczema. In particular, they have shorter fatty acids within their skin than unaffected individuals. This difference is suggested to lead to impaired skin barrier [70]. Atopic eczema is an inflammatory disease and thus processes of inflammation as discussed earlier will be active in the disease process [71]. As with other cases where a difference in fatty acid profiles has been established between individuals with disease and healthy individuals, it is not clear whether the fatty acid change causes the disease or is a response to disease, or possibly both, but it is a potential novel treatment route. Similar to people with atopic eczema, pruritic dogs have been shown to have a different fatty acid profile compared to dogs with healthy skin [72]. More recently, dogs with atopic dermatitis whose diets were supplemented with *n*-3 PUFA improved significantly more than those given the placebo [73]. As with human skin disease, it is not clear as to how this works, but it is an additional treatment option and

Cancer is the result of aberrant cellular processes. Many genes and proteins are differentially expressed in tumor tissue compared to nontumor tissue [74–77]. Thus, it is intuitive that fatty acid profiles are likely to be altered in tumors compared to nontumor tissue and this has

There have been studies showing that differential dietary intake of fatty acids can either reduce or increase risk of disease, including cancer. A meta-analysis of studies relating breast cancer risk with *n*-3 PUFA intake showed that overall increasing *n*-3 PUFA intake reduced the risk of developing breast cancer [78]. In transgenic mice in which males develop prostate cancer, *n*-3 PUFA intake from marine sources suppressed tumorigenesis [80]. This is also the case in people where there is reduced risk of developing prostate cancer with increased intake of marine *n*-3 PUFAs [81–83]. Longer chain *n*-3 PUFAs from non-marine sources, however, are

While ultimately work is required in whole organisms, cell lines are a valuable starting point for research. Of particular note in relation to veterinary medicine and fatty acids are two studies on canine tumor cell lines. The first is that of canine lymphoma cell lines; in this study, stearidonic acid was shown to sensitize cells to anticancer drugs, even when the cells

eases including heart disease and neurological dysfunction [55, 69].

**2.3. Fatty acids and skin disease**

182 Fatty Acids

area for further research.

**2.4. Cancer associations with fatty acids**

indeed been demonstrated in breast and prostate cancer [78, 79].

associated with an increased risk of prostate cancer [79, 82, 83].

Many animal and human studies have established that restriction of a range of nutrients within the maternal diet throughout pregnancy results in offspring that are programmed to be at increased risk of later hypertension and metabolic disease including diabetes and obesity [87–90]. This theory has become known as the "developmental origins of health and disease" (DOHaD) hypothesis. Fatty acid intake has been shown to have effects even before pregnancy as severe undernutrition of specific fatty acids has resulted in low reproductive rates in males and females. For example, in male cats, a linoleic deficient diet results in tubular degeneration of the testes and low fertility rates, and in females, the litters were not viable [91, 92].

Other studies have shown birth defects in offspring from females fed on low fatty acid diets but it also showed that arachidonate was a key contributor to viable offspring [93, 94]. In contrast, excess macronutrient intake has been implicated in the incidence of the metabolic syndrome is emerging in a number of rodent [95–97] and sheep studies [98]. Studies linking maternal over-nutrition to adverse offspring health in later life are conspicuously lacking, despite a huge effort in understanding the influence of maternal nutrition and its link to obesity. A number of rodent studies have established that a high-fat maternal diet leads to impaired offspring glucose and lipid metabolism [95–97, 99], but the influence of increasing other dietary components has not been investigated, perhaps due to the assumption that a high-fat or "junk food" diet is more prevalent in the western world. Rodent studies of increased fat intake during pregnancy are often associated with an overall decrease in food intake which limits their usefulness [97]. The timing of a nutritional insult is also important in determining the outcome for offspring, differential results have been observed in studies investigating early or late gestational nutritional insults in both animal [100, 101] and human studies [102]. As well as a high-fat diet increasing adipocyte and ectopic lipid accumulation, it may also decrease glycogen deposition in skeletal muscle. Increased plasma free fatty acids impair insulin-stimulated glucose disposal, including glycogenesis and glucose uptake—resulting in reduced skeletal muscle glycogen content [103]. Type-2 diabetes in humans is associated with a reduction in glycogen synthase and tissue glycogen [104], it is unknown whether a sub-optimal maternal diet will result in similar changes in offspring. Recent work has demonstrated that there are physiological [105–107] and emerging molecular differences between pigs with low, normal or high birth weights [108–111]. Extensive physiological examinations of low and high birth weight pigs, at 12 months of age showed that low birth weight pigs had increased fat depth and glucose intolerance and insulin resistance [105]. Also of interest is that, peroxisome proliferator-activated receptor (PPAR)α expression in skeletal muscle is positively correlated to birth weight in these pigs [110]. In younger pigs (7 or 14 days of postnatal age) designated low, normal or high to birth weight, molecular differences have been observed in adipose tissue and skeletal muscle genes known to regulate lipid metabolism including uncoupling proteins (UCPs), PPARα and γ, fatty acid-binding protein (FABP) 3 and 4 and the glucocorticoid receptor (GR) [108, 109, 111].

The role of PPARs is not just restricted to animals subjected to over-nutrition. Studies of maternal low protein diets in rats have demonstrated that post-weaning, offspring had significantly increased hepatic PPARα expression due to decreased methylation as a result of differences in overall dietary fat intake [112]. PPARs are a nuclear hormone receptor family that have attracted much interest due to their involvement in adipogenesis, lipid metabolism, insulin sensitivity, inflammation and blood pressure [113]. PPARγ regulates transcription of genes involved in lipid metabolism by binding to responsive elements in the promoters of respective genes. This transcription regulation stimulates fatty acid storage in adipose tissue by increasing the storage capacity and the quantity of fatty acids that enter adipocytes and also plays a key role in adipocyte differentiation, promoting the formation of mature lipid-laden adipocytes [114]. The activities of PPARγ are regulated by fatty acids (which are thought to be the endogenous ligands) [115]. PPARγ is often referred to as the "genetic sensor" for fat and a number of dietary studies have demonstrated an increase following high-fat feeding [116, 117], which may provide benefits to the animal by protecting against lipotoxic species [117]. PPARα also acts as a ligand-activated transcription factor and is expressed in tissues which have a high rate of fatty acid catabolism such as skeletal muscle and liver. The fibrate group of drugs has long been utilized as a synthetic ligand for PPARα, but endogenous ligands are still under investigation. Long-chain fatty acyl-CoAs and saturated fatty acids however are known to activate PPARα at micromolar ranges [118]. PPARα has a key role in stimulating lipid oxidation pathways to prevent storage of fats as well as increasing insulin sensitivity and glucose tolerance. The expression of PPARs may represent one of the molecular factors driving excess tissue lipid uptake, storage and production in animals that experienced a sub-optimal environment in utero, in particular low birth weight offspring; ectopic lipid storage, especially intramyocellular, is associated with glucose intolerance and type-2 diabetes [104, 119].

The regulation of fatty acids is also an important factor during the lactation period. A number of studies have shown that the relative fatty acid content of milk differs depending on the species. Donkeys have milk more similar to humans than cows, with lower levels of saturated fats and higher essential fatty acids than cows, more akin to humans [120, 121]. Milk, from humans, dog, and guinea pig are mostly comprised from long-chain fatty acids (48–54 acyl carbon atoms), cow, sheep, and goat, have more short-chain acids (28–54 acyl carbon atoms) and horses tended to have medium-chain fatty acids (26–54 carbon atoms range) [122]. Maternal diet can also have an impact on the fatty acid contents of her milk. This has been shown in many species from mice and sheep to humans [123–125]; the pregnancy status of the mother also vastly changes milk fatty acid composition [126]. These are important factors when assessing whether the mother is receiving an appropriate diet, assessing whether she is pregnant or not and whether milk replacement formulae contain the appropriate levels of fatty acids.

### **4. Fatty acid-binding proteins and lipid modulation**

tissue and skeletal muscle genes known to regulate lipid metabolism including uncoupling proteins (UCPs), PPARα and γ, fatty acid-binding protein (FABP) 3 and 4 and the glucocorti-

The role of PPARs is not just restricted to animals subjected to over-nutrition. Studies of maternal low protein diets in rats have demonstrated that post-weaning, offspring had significantly increased hepatic PPARα expression due to decreased methylation as a result of differences in overall dietary fat intake [112]. PPARs are a nuclear hormone receptor family that have attracted much interest due to their involvement in adipogenesis, lipid metabolism, insulin sensitivity, inflammation and blood pressure [113]. PPARγ regulates transcription of genes involved in lipid metabolism by binding to responsive elements in the promoters of respective genes. This transcription regulation stimulates fatty acid storage in adipose tissue by increasing the storage capacity and the quantity of fatty acids that enter adipocytes and also plays a key role in adipocyte differentiation, promoting the formation of mature lipid-laden adipocytes [114]. The activities of PPARγ are regulated by fatty acids (which are thought to be the endogenous ligands) [115]. PPARγ is often referred to as the "genetic sensor" for fat and a number of dietary studies have demonstrated an increase following high-fat feeding [116, 117], which may provide benefits to the animal by protecting against lipotoxic species [117]. PPARα also acts as a ligand-activated transcription factor and is expressed in tissues which have a high rate of fatty acid catabolism such as skeletal muscle and liver. The fibrate group of drugs has long been utilized as a synthetic ligand for PPARα, but endogenous ligands are still under investigation. Long-chain fatty acyl-CoAs and saturated fatty acids however are known to activate PPARα at micromolar ranges [118]. PPARα has a key role in stimulating lipid oxidation pathways to prevent storage of fats as well as increasing insulin sensitivity and glucose tolerance. The expression of PPARs may represent one of the molecular factors driving excess tissue lipid uptake, storage and production in animals that experienced a sub-optimal environment in utero, in particular low birth weight offspring; ectopic lipid storage, especially intramyocellular, is associated with glucose intolerance and

The regulation of fatty acids is also an important factor during the lactation period. A number of studies have shown that the relative fatty acid content of milk differs depending on the species. Donkeys have milk more similar to humans than cows, with lower levels of saturated fats and higher essential fatty acids than cows, more akin to humans [120, 121]. Milk, from humans, dog, and guinea pig are mostly comprised from long-chain fatty acids (48–54 acyl carbon atoms), cow, sheep, and goat, have more short-chain acids (28–54 acyl carbon atoms) and horses tended to have medium-chain fatty acids (26–54 carbon atoms range) [122]. Maternal diet can also have an impact on the fatty acid contents of her milk. This has been shown in many species from mice and sheep to humans [123–125]; the pregnancy status of the mother also vastly changes milk fatty acid composition [126]. These are important factors when assessing whether the mother is receiving an appropriate diet, assessing whether she is pregnant or not and whether milk replacement formulae contain the appro-

coid receptor (GR) [108, 109, 111].

184 Fatty Acids

type-2 diabetes [104, 119].

priate levels of fatty acids.

Fatty acids are now recognized as crucial components of cellular signaling cascades, in particular, those regulating lipid metabolism, as described above with PPARs. Research into fatty acids as signaling molecules is in its infancy, but it is well known that fatty acids are ligands for transcription factors. Fatty acids are carried through tissue membranes and in the cytosol by chaperones known as fatty acid-binding proteins (FABPs), of which there are a number of tissue-specific isoforms [127]. Knock-out mice not expressing the adipocyte-specific FABP4 exhibited protection from the metabolic effects (e.g. insulin resistance and hypercholesterolaemia) of a high-fat diet, suggesting FABP4 modulates a number of components of the metabolic syndrome [127]. In skeletal muscle, a fat-rich diet increases the expression of the cytosolic and plasma membrane specific FABP [128].

Insulin resistance is characterized by a decrease in the enzymes and proteins involved in lipid oxidation [129]. Lipogenesis and adipogenesis are modulated by the enzymes acetyl-CoA carboxylase 1 and 2 (ACC1 and ACC2, respectively) and AMP-activated protein kinase (AMPK); both enzymes are potential drug targets to treat obesity and the metabolic syndrome and AMPK has been suggested as a target for metformin [130, 131]. Briefly, ACC1 controls fatty acid biosynthesis and ACC2 controls fatty acid oxidation. ACC1 catalyses the conversion of acetyl-COA to malonyl-CoA, therefore modulating the rate limiting step of long-chain fatty acid biosynthesis in adipose tissue. ACC2 is expressed in skeletal muscle, where the product malonyl-CoA inhibits fatty acid oxidation. The AMPKα subunit is activated during periods of metabolic stress (e.g. increased AMP/ATP ratio) by phosphorylation and inhibits the activity of ACC1 and 2, thus promoting fatty acid oxidation, glucose uptake and inhibits lipid synthesis [132] and thereby reducing ectopic lipid storage. An isocaloric high-fat diet has been shown to inhibit AMPK in rats [133]. Despite great potential for modulation by maternal diet, there are few DOHAD studies of ACC and AMPK expression; however, early studies of an obese pregnant ewe model have shown decreased AMPK signaling in fetal offspring muscle [98].

### **5. Future fatty acid research and medicine**

Although artificially induced disease often only replicates a small aspect of disease and does not reflect the typically longer time scales involved in natural disease progression in both humans and animals [134, 135], these studies can be valuable when compared to naturally occurring diseases in order to understand mechanisms and development. All of the 'natural population' studies discussed in this chapter may have their own caveats too. Differences in diet, age, sex and even pre-clinical symptoms and diagnosis can all affect the results observed in both disease and fatty acid states. This chapter has concentrated on development, cardiovascular disease, cancer and immunity but differing fatty acids have been implicated or associated with in a number of diseases and disorders ranging from human, rodent and canine epilepsy through to canine ADHD and reproductive ability [92, 136, 137].

Fatty acid profiling has important potential applications as a diagnosis tool across the species, especially in cases where pre-clinical symptoms are difficult to observe. Although it is not always necessarily known if differences in fatty acid profiles are contributing to the initiation of disease or are a response to disease processes, these differences could be drug targets [26, 138–140]. In addition, there are genes that contribute to fatty acid profile composition and if a particular part of the pathway is shown to be different in individuals with disease compared to healthy individuals, these could be likely genes for candidate gene studies in the future [141, 142]. The scientific methodologies available for looking at lipid levels have also progressed over the years; just one example is the use of proton magnetic resonance spectroscopy of protons (H-MRS) to assess cardiac lipids in a non-invasive manner [52]. This is a valuable tool for animal health and welfare, and there are additional uses in looking at metabolism and fatty acids. Much of the present work involves looking at genes and lipid levels of animals intended for the meat industry. An example is the evidence that differing polymorphisms in genes can result in differing meat quality traits. This includes fatty acid synthase (FASN) which was found to correlate with meat weight loss during the first salting of dry-cured ham production [143], meat quality including marbling in cattle [144] and playing a role in the mammary gland and milk in goats and cattle [145, 146], in addition to many other roles. Differing H-FABP polymorphisms/expression levels have also been related to growth rate and size of beef cattle and chickens and could therefore provide useful markers for breeding [147, 148].

Research into the links between fatty acids and differing developmental stages and disease states is increasing in both humans and animals and provides the potential for innovative diagnostic and treatments tools.

### **Acknowledegments**

This work was supported by the Biotechnology and Biological Sciences Research Council [grant number BB/J014508/1], by generous funding to Catrin S. Rutland from the BBSRC University of Nottingham Doctoral Training Programme.

### **Author details**

Siobhan Simpson1† , Alison Mostyn1,2† and Catrin S. Rutland<sup>1</sup> \*

\*Address all correspondence to: catrin.rutland@nottingham.ac.uk

1 Faculty of Medicine, School of Veterinary Medicine and Science, University of Nottingham, UK

2 Faculty of Medicine, School of Health Sciences, University of Nottingham, UK

† Joint first authors

### **References**

Fatty acid profiling has important potential applications as a diagnosis tool across the species, especially in cases where pre-clinical symptoms are difficult to observe. Although it is not always necessarily known if differences in fatty acid profiles are contributing to the initiation of disease or are a response to disease processes, these differences could be drug targets [26, 138–140]. In addition, there are genes that contribute to fatty acid profile composition and if a particular part of the pathway is shown to be different in individuals with disease compared to healthy individuals, these could be likely genes for candidate gene studies in the future [141, 142]. The scientific methodologies available for looking at lipid levels have also progressed over the years; just one example is the use of proton magnetic resonance spectroscopy of protons (H-MRS) to assess cardiac lipids in a non-invasive manner [52]. This is a valuable tool for animal health and welfare, and there are additional uses in looking at metabolism and fatty acids. Much of the present work involves looking at genes and lipid levels of animals intended for the meat industry. An example is the evidence that differing polymorphisms in genes can result in differing meat quality traits. This includes fatty acid synthase (FASN) which was found to correlate with meat weight loss during the first salting of dry-cured ham production [143], meat quality including marbling in cattle [144] and playing a role in the mammary gland and milk in goats and cattle [145, 146], in addition to many other roles. Differing H-FABP polymorphisms/expression levels have also been related to growth rate and size of beef cattle and chickens and could therefore provide useful

Research into the links between fatty acids and differing developmental stages and disease states is increasing in both humans and animals and provides the potential for innovative

This work was supported by the Biotechnology and Biological Sciences Research Council [grant number BB/J014508/1], by generous funding to Catrin S. Rutland from the BBSRC

and Catrin S. Rutland<sup>1</sup>

1 Faculty of Medicine, School of Veterinary Medicine and Science, University of Nottingham,

2 Faculty of Medicine, School of Health Sciences, University of Nottingham, UK

\*

markers for breeding [147, 148].

diagnostic and treatments tools.

University of Nottingham Doctoral Training Programme.

, Alison Mostyn1,2†

\*Address all correspondence to: catrin.rutland@nottingham.ac.uk

**Acknowledegments**

**Author details**

Siobhan Simpson1†

† Joint first authors

UK

186 Fatty Acids


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