Analytical Tools for Lipid Assessment in Biological Assays

*Banny Silva Barbosa Correia, Raquel Susana Torrinhas, William Yutaka Ohashi and Ljubica Tasic*

## **Abstract**

Lipids are heterogeneous biological molecules with many important roles. In human body, lipids can be energy substrates, steroid hormones, inflammatory lipid mediators, transporters, and feature as structural cell and organelle membrane elements. At the cell membrane, lipids influence the distribution of surface proteins and, in part, protein signaling and, consequently, the activation of transcriptional factors. One of the best explored relationships in chemistry and science is the structure/activity one. Therefore, if the composition of a mixture is discovered and the structure of its components is known, a task of proposing relationship among all components and their activity would be closer to understanding. There are many powerful and advantageous analytical and bioanalytical tools available for the study of lipids, but all show at least some limitations. Knowing the advantages/disadvantages of each technique is essential for choosing the most appropriate one for the analysis as to answer a scientific question about lipid composition and role within a biological model. Often, inexperience and little familiarity with the cited analytical resources may limit the validity of the obtained results. Our chapter aims to present and discuss different tools available for the study of lipids and their main applications in biological assays.

**Keywords:** lipids, bioanalytical tools, gas chromatography, mass spectrometry, nuclear magnetic resonance

## **1. Introduction**

Lipids are a very heterogeneous group of biological molecules. Some of the most studied lipids are built from the fatty acids (FAs) or isoprenyl groups. FAs are carboxylic acids composed by an even number of carbon atoms connected by single or double bonds with a methyl group end. FAs can be classified into very long (>20 carbons), long (14–20 carbons), medium (6–12 carbons), and short (up to 6 carbons)-chain FAs, as well as saturated (no double chains), monounsaturated (1 double bond), and polyunsaturated (PUFAs, >1 double bond) FAs. Furthermore, unsaturated fatty acids can receive its omega (n) assignment according to the first double-bond position from the end methyl group. Biosynthetically, endogenous FAs have been made from acetyl-CoA/malonyl-CoA [1–3].

FAs represent a class of lipids on their own and do not make part of all lipids [4]. Some lipids, which are not formed from FAs but are biosynthetically related to them, are the polyketides, formed from the acetyl units. Other unsaponifiable lipids are built from isoprene units, molecules with five carbons with a branch structure and alternated double bonds. Isoprenes have their biosynthesis in mevalonate (vegetables) or deoxyxylulose phosphate (animals) pathways. They can form sterols and prenols [2]; some sterols can also have FAs in their structure [3].

Actually, lipids comprise eight main classes within different chemical characteristics: fatty acids (1), glycerolipids (2), glycerophospholipids (3), sphingolipids (4), sterols (5), prenols (6), saccharolipids (7), and polyketides (8) (**Figure 1**) [3]. These classes show a high diversity of molecules and are grouped into several subclasses. Lipid classification based on their chemical information, described by the headgroup and the type of a linkage between the head group and aliphatic chains [5, 6] is the most used among biochemists. Investigators have estimated the presence of ~180,000 lipid species in nature and ~40 common fatty acids as building blocks [7]. At the moment, 43,109 structurally distinct lipids are already registered at the Lipid MAPS consortium.

The high diversity of lipids reflects their multiple biological functions and can be attributed to the wide variety of their building blocks and numerous possible permutations [6, 8]. In the human body, lipids serve as: substrates for the synthesis of energy (9.3 kcal/g), steroid hormones, inflammatory lipid mediators, vitamins or liposoluble vitamins transportation, and structural elements of cell and organelle membranes [9–11]. As a part of the cell membrane, lipids can influence the distribution of surface proteins, protein signaling (as part of lipid rafts or as second messengers), and consequently, the activation of transcriptional factors [12, 13]. This means that besides their recognized biological functions, lipids can influence protein signaling and synthesis.

#### **Figure 1.**

*Biosynthetic lipid network. Acetyl-CoA: fatty acids-FAs (class 1) are synthetized, enabling the production of other lipid classes: 2 (glycerolipids-GLs), 3 (glycerophospholipids-GPs), 4 (sphingolipids-SPs), and 7 (saccharolipids-SLs), as well the class of eicosanoids. Acetyl-CoA can also generate the class 8 (polyketides-PKs) and isopentenyl diphosphate molecule, through mevalonate. On the other side, isoprenyl is used as starting substrate for producing lipid classes 6 (prenols-PRs) and 5 (sterols-STs). Figure was inspired on Quhenberger et al. [4].*

**23**

*(C24:5 n-3), and tetracosahexaenoic acid (C24:6 n-3).*

**Figure 2.**

*Analytical Tools for Lipid Assessment in Biological Assays*

In a cell, lipids show different compositions, tens of thousands to hundreds of thousands of compounds, and concentrations from a mol/mg to nmol/mg of protein [5]. Facing the biological relevance of lipids, it is not surprising that the human organism has sophisticated machineries for the FA synthesis when its dietary supply flaws. Saturated and monounsaturated FAs can be endogenously generated from glucose and amino acids through enzymatic elongation (by adding units of two carbons) and desaturation (by forming new double bonds) reactions. However, a pitiful lack of the desaturating enzymes ∆-12 and ∆-15 desaturases preclude humans to add double bonds before the ninth carbon at the end of the methyl extremity for the synthesis of the polyunsaturated fatty acids (PUFAs) n-linoleic acid (C18:2 n-6, LA) and alpha-linolenic acid (C18:3 n-3, ALA). Consequently, LA and ALA are obtained exclusively from diet and, then, called as essentials. After ingestion, LA and ALA compete for sequential enzymatic processes of elongation and desaturation until their conversion into longer chain PUFAs: arachidonic acid (C20:4 n-6, ARA) from LA and eicosapentaenoic acid (C20:5 n-3, EPA) or docosahexaenoic acid

ARA, EPA, and DHA have a high clinical interest once they influence the composition and steady-state of cell membranes. Also, they are precursors of the lipid mediators named eicosanoids involved in the activation of the inflammatory

*Synthesis of lipid mediators from eicosapentaenoic (C20:2 n-6, EPA), docosahexaenoic (C22:6 n-3, DHA), and arachidonic (C20:4 n-6, ARA) acids. EPA, DHA, and ARA are previously synthetized from n-3 and n-6 fatty acid families in reactions mediated by enzymes: 1—desaturase, 2—elongase, 3—peroxisomal fatty acyl-CoA oxidase, 4—lipoxygenase (LOX), and 5—cyclooxygenase (COX). The cellular bioavailability of EPA decreases the production of ARA-produced eicosanoids, which include prostaglandins (PG)E2, thromboxane (TX) A2, and leukotriene (LT)B4. These eicosanoids have a higher pro-inflammatory potential than those contra parts produced from EPA (PGE5, TXA3, and LTB5) in promoting vasodilation and leukocyte chemotaxis and adhesion, events that stimulate the migration of neutrophils into the damaged tissue. As part of the neutrophil-monocyte sequence of inflammation, eicosanoids are no longer produced to initiate the synthesis of resolvins, protectins and maresins, lipid mediators from EPA and DHA. Other fatty acids shown are: linoleic acid (C18:2 n-6, LA), gamma-linolenic acid (C18:3 n-6, GLA), dihomo-gamma-linolenic acid (C20:3 n-6, DGLA), adrenic acid (C22:4 n-6), tetracosatetraenoic acid (C24:4 n-6), tetracosapentaenoic acid (C24:5 n-6), docosapentaenoic acid (C22:5 n-6), oleic acid (C18:1 n-9), octadecadienoic acid (C18:2 n-9), alphalinolenic acid (C18:3 n-3, ALA), stearidonic acid (C18:4 n-3, SDA), eicosatrienoic acid (C20:3 n-3, ETE), eicosatetraenoic acid (C20:4 n-3, ETA), docosapentaenoic acid (C22:5 n-3, DPA), tetracosapentaenoic acid* 

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

(C22:6 n-3, DHA) from ALA [14].

*Analytical Tools for Lipid Assessment in Biological Assays DOI: http://dx.doi.org/10.5772/intechopen.81523*

*Advances in Lipid Metabolism*

MAPS consortium.

protein signaling and synthesis.

are built from isoprene units, molecules with five carbons with a branch structure and alternated double bonds. Isoprenes have their biosynthesis in mevalonate (vegetables) or deoxyxylulose phosphate (animals) pathways. They can form sterols and

Actually, lipids comprise eight main classes within different chemical characteristics: fatty acids (1), glycerolipids (2), glycerophospholipids (3), sphingolipids (4), sterols (5), prenols (6), saccharolipids (7), and polyketides (8) (**Figure 1**) [3]. These classes show a high diversity of molecules and are grouped into several subclasses. Lipid classification based on their chemical information, described by the headgroup and the type of a linkage between the head group and aliphatic chains [5, 6] is the most used among biochemists. Investigators have estimated the presence of ~180,000 lipid species in nature and ~40 common fatty acids as building blocks [7]. At the moment, 43,109 structurally distinct lipids are already registered at the Lipid

The high diversity of lipids reflects their multiple biological functions and can be attributed to the wide variety of their building blocks and numerous possible permutations [6, 8]. In the human body, lipids serve as: substrates for the synthesis of energy (9.3 kcal/g), steroid hormones, inflammatory lipid mediators, vitamins or liposoluble vitamins transportation, and structural elements of cell and organelle membranes [9–11]. As a part of the cell membrane, lipids can influence the distribution of surface proteins, protein signaling (as part of lipid rafts or as second messengers), and consequently, the activation of transcriptional factors [12, 13]. This means that besides their recognized biological functions, lipids can influence

*Biosynthetic lipid network. Acetyl-CoA: fatty acids-FAs (class 1) are synthetized, enabling the production of other lipid classes: 2 (glycerolipids-GLs), 3 (glycerophospholipids-GPs), 4 (sphingolipids-SPs), and 7 (saccharolipids-SLs), as well the class of eicosanoids. Acetyl-CoA can also generate the class 8 (polyketides-PKs) and isopentenyl diphosphate molecule, through mevalonate. On the other side, isoprenyl is used as starting substrate for producing lipid classes 6 (prenols-PRs) and 5 (sterols-STs). Figure was inspired on Quhenberger* 

prenols [2]; some sterols can also have FAs in their structure [3].

**22**

**Figure 1.**

*et al. [4].*

In a cell, lipids show different compositions, tens of thousands to hundreds of thousands of compounds, and concentrations from a mol/mg to nmol/mg of protein [5]. Facing the biological relevance of lipids, it is not surprising that the human organism has sophisticated machineries for the FA synthesis when its dietary supply flaws. Saturated and monounsaturated FAs can be endogenously generated from glucose and amino acids through enzymatic elongation (by adding units of two carbons) and desaturation (by forming new double bonds) reactions. However, a pitiful lack of the desaturating enzymes ∆-12 and ∆-15 desaturases preclude humans to add double bonds before the ninth carbon at the end of the methyl extremity for the synthesis of the polyunsaturated fatty acids (PUFAs) n-linoleic acid (C18:2 n-6, LA) and alpha-linolenic acid (C18:3 n-3, ALA). Consequently, LA and ALA are obtained exclusively from diet and, then, called as essentials. After ingestion, LA and ALA compete for sequential enzymatic processes of elongation and desaturation until their conversion into longer chain PUFAs: arachidonic acid (C20:4 n-6, ARA) from LA and eicosapentaenoic acid (C20:5 n-3, EPA) or docosahexaenoic acid (C22:6 n-3, DHA) from ALA [14].

ARA, EPA, and DHA have a high clinical interest once they influence the composition and steady-state of cell membranes. Also, they are precursors of the lipid mediators named eicosanoids involved in the activation of the inflammatory

#### **Figure 2.**

*Synthesis of lipid mediators from eicosapentaenoic (C20:2 n-6, EPA), docosahexaenoic (C22:6 n-3, DHA), and arachidonic (C20:4 n-6, ARA) acids. EPA, DHA, and ARA are previously synthetized from n-3 and n-6 fatty acid families in reactions mediated by enzymes: 1—desaturase, 2—elongase, 3—peroxisomal fatty acyl-CoA oxidase, 4—lipoxygenase (LOX), and 5—cyclooxygenase (COX). The cellular bioavailability of EPA decreases the production of ARA-produced eicosanoids, which include prostaglandins (PG)E2, thromboxane (TX) A2, and leukotriene (LT)B4. These eicosanoids have a higher pro-inflammatory potential than those contra parts produced from EPA (PGE5, TXA3, and LTB5) in promoting vasodilation and leukocyte chemotaxis and adhesion, events that stimulate the migration of neutrophils into the damaged tissue. As part of the neutrophil-monocyte sequence of inflammation, eicosanoids are no longer produced to initiate the synthesis of resolvins, protectins and maresins, lipid mediators from EPA and DHA. Other fatty acids shown are: linoleic acid (C18:2 n-6, LA), gamma-linolenic acid (C18:3 n-6, GLA), dihomo-gamma-linolenic acid (C20:3 n-6, DGLA), adrenic acid (C22:4 n-6), tetracosatetraenoic acid (C24:4 n-6), tetracosapentaenoic acid (C24:5 n-6), docosapentaenoic acid (C22:5 n-6), oleic acid (C18:1 n-9), octadecadienoic acid (C18:2 n-9), alphalinolenic acid (C18:3 n-3, ALA), stearidonic acid (C18:4 n-3, SDA), eicosatrienoic acid (C20:3 n-3, ETE), eicosatetraenoic acid (C20:4 n-3, ETA), docosapentaenoic acid (C22:5 n-3, DPA), tetracosapentaenoic acid (C24:5 n-3), and tetracosahexaenoic acid (C24:6 n-3).*

response. While ARA is a precursor of pro-inflammatory, immunosuppressive, and pro-thrombotic eicosanoids, EPA competes with ARA for lipoxygenase (LOX) and cyclooxygenase (COX) enzymes to generate functionally less intense and antithrombotic mediators [10]. Furthermore, EPA and DHA are precursors of resolvins and DHA is a precursor of protectins and maresins. These lipid mediators are collectively called as specialized pro-resolving mediators and have a relevant role in the inflammation resolution and homeostasis restoring [15]. In conjunction, these observations traduce an anti-inflammatory and pro-resolving potential of EPA and DHA (**Figure 2**).

Moreover, EPA, DHA, and their metabolites can exert anti-inflammatory and metabolic effects by modulating the activity of transcriptional factors, such as nuclear kappa B factor (NFκB), nuclear factor E2-related factor 2 (Nfr2), peroxisome proliferator-activated receptor (PPAR), and sterol regulatory elementbinding proteins (SREBP). Due to their abilities, EPA and DHA can influence the transcription of genes enrolled in inflammation, cell survival, oxidative stress, and in carbohydrate and lipid metabolism [16]. Some of the EPA and DHA functions arise from the capacity of these n-3 PUFAs (mainly DHA) to interfere in protein receptors signaling by disrupting lipid rafts, membrane microdomains rich in saturated FAs (mainly cholesterol) who confer a rigidity needed for some protein dimerization through the fluid cell membrane [17, 18].

Due to biological properties, the importance of EPA and DHA for human health has been highly discussed and investigated by basic, translational, and epidemiologic scientists. However, studies on lipids and their biological relevance are not limited to n-3 PUFAs or other individual lipids, but also include the analysis of all

#### **Figure 3.**

*The most common analytical techniques used in analyses of lipids and lipidomes are gas chromatography (GC), mass spectrometry (MS), and nuclear magnetic resonance (NMR) spectroscopy. These techniques show some vantages and weaknesses and could be used in combination with other techniques in so-called hyphenated bioanalytical methods. All enable qualitative and quantitative analyses of lipids, but GC needs additional step in sample preparation as to increase the volatility of the compounds; thus, not all lipids could be analyzed by GC. Also, GC requires greater sample quantities when compared to MS, which is most sensitive. MS analyses require the use of ionization techniques, such as electron and chemical ones for gas samples, while electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) are usually applied for liquid and solid samples. NMR is the only nondestructive technique and allows the noninvasive lipid analysis in intact cells and tissues, and enables to investigate changes in lipid and dynamic structures in biochemical cell functionalization, but it is not sufficiently sensitive and universal when compared to MS.*

**25**

*Analytical Tools for Lipid Assessment in Biological Assays*

hypothesis, with focus on nutrition and metabolism aspects.

point until they get to an electronic detector [20].

**2. Gas chromatography: principles, strengths, and weaknesses**

lipid species from a biological sample—the lipidome. Because lipids are intermediates and even signaling molecules of metabolic pathways, the lipidomic response (change of the lipidome pattern of a biological sample) to nutritional, pharmacological, or any intervention (i.e., surgery, exercise) treatments can reflect their biological effects [5]. Studies on lipidome can also add to the knowledge on the lipid content of a nutritional source (i.e., fish) aiming to found ones with the high n-3 PUFAs, for instance. These are examples of many applications of lipid analysis in

There are several tools available for the study of individual lipids and lipidome (the total lipid content in a cell or an organism), all with their advantages and limitations (**Figure 3**). Understanding these points is essential for the application of that most appropriate techniques to answer a scientific question on lipids within a biological model. Often little familiarity with these analytical resources may limit the validity of the results. This chapter aims to present and discuss different tools available for different applications in the study of lipids aiming to assess biological

According to the principles of chromatographic techniques, the gas chromatography (GC) is applied when aimed to separate organic compounds from a mixture in the gas form. For this purpose, the GC uses interaction among the sample components and the stationary phase and the mobile gas phase. After lipid extraction, the samples (lipid mixture) are usually liquids and must be exposed to a high temperature at the gas chromatograph entrance (injector). Vaporized, the samples are carried by a gas, which is usually a nonheavy and inert gas (i.e., hydrogen, helium), through a long capillary column containing a high or low polarity material

The gaseous compounds generated from the vaporized sample interact with the stationary phase what allows each compound to elute/separate at a different time (retention time). Because GC considers both chemical and physical properties of the vaporized compounds, those with more chemical affinity to the stationary phase will take longer time to be removed from the column and the temperature will influence the overall process. This explains why the column stays in an oven, which is programmed to work at different temperature ranges (i.e., temperature programming) in which the compounds are carried out by the gas according to their boiling

At the end of GC analysis, the electronic detector generates a chromatogram based on retention time by intensity. This allows a qualitative identification of the lipid compounds by comparing their retention times with certified standard using the flame ionization detector (FID) or by deduction of spectra information using a mass spectrometer as detector. Lipid quantification can also be performed using analytical procedures of external or internal certified standard in GC analysis [21]. Main points to be considered when assessing FAs by GC analysis are the carrier gas flow rate, column length, and the temperature because these can influence the order or retention time of the lipid compounds and then must be precisely standardized [22]. The column length of the stationary phase influences the resolution of the analytes, once the number of theoretical plates (hypothetical zone in which two phases establish an equilibrium with each other) is respectively high in longer column. As fat and oils have high boiling points not supported by the stationary phase, a previous derivatization reaction step is required after lipid extraction from the biological sample, in which triacylglycerol and free fatty acids are transformed

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

biological systems.

(stationary phase) [19].

#### *Analytical Tools for Lipid Assessment in Biological Assays DOI: http://dx.doi.org/10.5772/intechopen.81523*

*Advances in Lipid Metabolism*

DHA (**Figure 2**).

**24**

**Figure 3.**

*The most common analytical techniques used in analyses of lipids and lipidomes are gas chromatography (GC), mass spectrometry (MS), and nuclear magnetic resonance (NMR) spectroscopy. These techniques show some vantages and weaknesses and could be used in combination with other techniques in so-called hyphenated bioanalytical methods. All enable qualitative and quantitative analyses of lipids, but GC needs additional step in sample preparation as to increase the volatility of the compounds; thus, not all lipids could be analyzed by GC. Also, GC requires greater sample quantities when compared to MS, which is most sensitive. MS analyses require the use of ionization techniques, such as electron and chemical ones for gas samples, while electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) are usually applied for liquid and solid samples. NMR is the only nondestructive technique and allows the noninvasive lipid analysis in intact cells and tissues, and enables to investigate changes in lipid and dynamic structures in biochemical cell* 

response. While ARA is a precursor of pro-inflammatory, immunosuppressive, and pro-thrombotic eicosanoids, EPA competes with ARA for lipoxygenase (LOX) and cyclooxygenase (COX) enzymes to generate functionally less intense and antithrombotic mediators [10]. Furthermore, EPA and DHA are precursors of resolvins and DHA is a precursor of protectins and maresins. These lipid mediators are collectively called as specialized pro-resolving mediators and have a relevant role in the inflammation resolution and homeostasis restoring [15]. In conjunction, these observations traduce an anti-inflammatory and pro-resolving potential of EPA and

Moreover, EPA, DHA, and their metabolites can exert anti-inflammatory and metabolic effects by modulating the activity of transcriptional factors, such as nuclear kappa B factor (NFκB), nuclear factor E2-related factor 2 (Nfr2), peroxisome proliferator-activated receptor (PPAR), and sterol regulatory elementbinding proteins (SREBP). Due to their abilities, EPA and DHA can influence the transcription of genes enrolled in inflammation, cell survival, oxidative stress, and in carbohydrate and lipid metabolism [16]. Some of the EPA and DHA functions arise from the capacity of these n-3 PUFAs (mainly DHA) to interfere in protein receptors signaling by disrupting lipid rafts, membrane microdomains rich in saturated FAs (mainly cholesterol) who confer a rigidity needed for some protein

Due to biological properties, the importance of EPA and DHA for human health has been highly discussed and investigated by basic, translational, and epidemiologic scientists. However, studies on lipids and their biological relevance are not limited to n-3 PUFAs or other individual lipids, but also include the analysis of all

dimerization through the fluid cell membrane [17, 18].

*functionalization, but it is not sufficiently sensitive and universal when compared to MS.*

lipid species from a biological sample—the lipidome. Because lipids are intermediates and even signaling molecules of metabolic pathways, the lipidomic response (change of the lipidome pattern of a biological sample) to nutritional, pharmacological, or any intervention (i.e., surgery, exercise) treatments can reflect their biological effects [5]. Studies on lipidome can also add to the knowledge on the lipid content of a nutritional source (i.e., fish) aiming to found ones with the high n-3 PUFAs, for instance. These are examples of many applications of lipid analysis in biological systems.

There are several tools available for the study of individual lipids and lipidome (the total lipid content in a cell or an organism), all with their advantages and limitations (**Figure 3**). Understanding these points is essential for the application of that most appropriate techniques to answer a scientific question on lipids within a biological model. Often little familiarity with these analytical resources may limit the validity of the results. This chapter aims to present and discuss different tools available for different applications in the study of lipids aiming to assess biological hypothesis, with focus on nutrition and metabolism aspects.

## **2. Gas chromatography: principles, strengths, and weaknesses**

According to the principles of chromatographic techniques, the gas chromatography (GC) is applied when aimed to separate organic compounds from a mixture in the gas form. For this purpose, the GC uses interaction among the sample components and the stationary phase and the mobile gas phase. After lipid extraction, the samples (lipid mixture) are usually liquids and must be exposed to a high temperature at the gas chromatograph entrance (injector). Vaporized, the samples are carried by a gas, which is usually a nonheavy and inert gas (i.e., hydrogen, helium), through a long capillary column containing a high or low polarity material (stationary phase) [19].

The gaseous compounds generated from the vaporized sample interact with the stationary phase what allows each compound to elute/separate at a different time (retention time). Because GC considers both chemical and physical properties of the vaporized compounds, those with more chemical affinity to the stationary phase will take longer time to be removed from the column and the temperature will influence the overall process. This explains why the column stays in an oven, which is programmed to work at different temperature ranges (i.e., temperature programming) in which the compounds are carried out by the gas according to their boiling point until they get to an electronic detector [20].

At the end of GC analysis, the electronic detector generates a chromatogram based on retention time by intensity. This allows a qualitative identification of the lipid compounds by comparing their retention times with certified standard using the flame ionization detector (FID) or by deduction of spectra information using a mass spectrometer as detector. Lipid quantification can also be performed using analytical procedures of external or internal certified standard in GC analysis [21].

Main points to be considered when assessing FAs by GC analysis are the carrier gas flow rate, column length, and the temperature because these can influence the order or retention time of the lipid compounds and then must be precisely standardized [22]. The column length of the stationary phase influences the resolution of the analytes, once the number of theoretical plates (hypothetical zone in which two phases establish an equilibrium with each other) is respectively high in longer column. As fat and oils have high boiling points not supported by the stationary phase, a previous derivatization reaction step is required after lipid extraction from the biological sample, in which triacylglycerol and free fatty acids are transformed

into their respective free fatty esters with lower boiling points (transesterification/ esterification reaction) [23]. Several methods are available for FAs derivatization [24], and the most applied ones are described in the 969.33 AOAC's method [25].

Particularly for cholesterol analysis, the samples preparation must consider a derivatization reaction. This allows to block protic sites of steroids obtained after an unsaponifiable lipid extraction [26] had been performed, and also, to diminish dipole-dipole interactions, to increase the volatility of the compounds, and to generate products with reduced polarity. Cholesterol derivatization is usually achieved by using trimethylsilyl compounds as reagents (silylation reaction). A common method for this purpose is described by Bowden and collaborators [27], in which *N*,*O*-bis(trimethylsilyl-trifluoroacetamide/trimethylchlorosilane)—BSTFA/ TMCS is used.

Nowadays, other more modern analytical tools than GC (next-generation techniques) do not require sample derivatization for lipid analysis. Needed lipid derivatization can be then consider a quite limitation step of the technique. In comparison with next-generation techniques, GC also implies in using greater sample quantities. This may be the main limitation in biological assays, which usually lead with restricted sample amounts. Nevertheless, by using certified standard and a powerful detector as FID, GC has the advantage to allow a precise and complete (by burning every compound, no one is lost in the detection) quantification of lipid compounds from biological samples, not always achieved by the other analytical techniques. In this context, GC continues to be accepted as an efficient and simple technique for FA and sterol analyses, mainly when combined with mass spectrometry (MS, detailed later in this chapter).

#### **2.1 Gas chromatography: application in biological assays**

In biological issues, GC is largely applied to assess the FA and cholesterol contents in animal models or human fluids and tissues, as biological markers of FA ingestion and cell incorporation. The technique is a powerful tool in studies assessing the effect of FA supplementation on a specific biologic response. For instance, the endogenous synthesis of EPA and DHA from ALA is low in humans, who have in the ingestion, oily fishes as the most relevant source. Therefore, studies on n-3 PUFAs have been focused on the effect of fish ingestion or fish oil/EPA/DHA supplementation in several clinical conditions, and cell and disease models. In such studies, the treatment compliance or effectiveness can be reflected by the cell or circulate contends of n-3 PUFAs [28]. Furthermore, GC can be applied to validate data generated by other lipidomic techniques.

A practical example in using GC for treatment compliance is the study of Nogueira et al. [29] assessing the effect of n-3 PUFAs supplementation in patients with nonalcoholic steatohepatitis against placebo (mineral oil). In this study, GC highlighted a similar increase in n-3 PUFAs plasma in both n-3 PUFAs- and placebo-treated patients. Because the authors have controlled compliance of n-3 PUFAs, they were able to discover off-protocol intake of PUFAs by some patients from the placebo group. When studying biochemical markers of lipid intake and cell incorporation, the biological sample nature matters. For instance, plasma and red blood samples can reflect periods of weeks and months of FAs ingestion and their effects, respectively, while the adipose tissue is the reference method, once it reflects such variables for years [28].

The study of Ravacci et al. [30] can illustrate the use of GC to assess treatment compliance. Applying this technique, the authors were able to demonstrate that the treatment of a lineage of breast cancer cell overexpressing HER-2 with DHA increased its availability in the cell membrane and was associated with the

**27**

*Analytical Tools for Lipid Assessment in Biological Assays*

disruption of surface lipid rafts that sustained cell signal for survival. Regarding the use of GC for data validation, a practical example is the study of Ouldamer et al. [31], which applied the technique to validate the fatty acid information generated

**3. Mass spectrometry: principles, strengths, and weaknesses**

called as genomics, transcriptomics, and proteomics, respectively [32].

tion/ionization (MALDI) are usually applied [33].

signal-to-noise ratio and reproducibility [32].

centration units, goes from a mol L<sup>−</sup><sup>1</sup>

and the abundances of each ion present in a sample are reported.

system, as GC (early mentioned) and liquid chromatography (LC) [35]. Data obtained by MS are displayed as spectra of the relative abundance of detected ions as a function of the corresponding m/z. By correlating the known masses (e.g., an entire molecule) to the identified masses, or through the

H NMR analysis on the PUFAs n-3 DHA and EPA content in the adipose tissue of mammary tumor model in rats exposed to controlled dietary intake of lipids.

Modernization of MS used for lipid analysis raised the concept of lipidomics. Lipidomics is an emerging science that aims to analyze the total lipid content found in a cell or tissue (lipidome) through the application of analytical chemistry principles and techniques. As a part of the omics sciences, the processes applied in the lipidomic analysis are analogous to those applied in other life-building macromolecules, such as deoxyribonucleic acid (DNA), ribonucleic acid (RNA), and proteins,

The basic principle of the MS technique is founded on the detection of the abundance of ions by their mass/charge ratio (m/z). To allow the analysis, such ions of compounds are generated by suitable methods and ions are separated according to their m/z. Ionization techniques can break some sample's molecules into charged fragments and are chosen according to the physical state of the sample. Also, the efficiency of various ionization mechanisms for the unknown species might help when picking the most appropriate ionization technique. The most common ones for gaseous samples are the electron and chemical ionizations, while for liquid and solid samples, the electrospray ionization (ESI) and matrix-assisted laser desorp-

Advances in ESI-MS and MALDI-MS have greatly facilitated lipidomic analysis [34] and enabled a great progress in lipid metabolic discoveries. This is because ESI is one of the softest ionization techniques, in which some complex dimers and solvent adducts can also be detected at the end. The efficiency of lipid ionization in ESI is incomparably higher than achieved by other traditional MS ion sources. MALDI-MS counts on a good solubility of analytes (lipids) and a matrix (for example, 2,5-dihydrobenzoic acid) in organic solvents, and provides excellent

Mass spectrometers are made from three components: the ion source (1), which converts a sample into ions that are targeted through the mass analyzer (2) and run into the detector (3). The mass analyzer acts as ions organizer (classifier) using ion m/z ratios. This component accelerates ions as they face a strong electromagnetic field. The detector measures charged particles, such as an electron multiplier [35],

An advantage of MS is its high sensibility. A detection limit, expressed in con-

as the instruments modernize. For example, the instrument response factor for any individual molecular species detected is essentially identical within experimental error after 13C deisotoping if the analysis is performed properly [34]. Also, MS ion source can act as a separation device if set to selectively ionize just a certain lipid class. Thus, it is feasible to analyze different classes of lipids and individual molecular species with high efficiency without prior chromatographic separation. Nevertheless, depending on the analysis aims, MS can also be combined with a chromatography

to as low as fmol L<sup>−</sup><sup>1</sup>

and surely shall improve

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

by the 1

*Advances in Lipid Metabolism*

TMCS is used.

etry (MS, detailed later in this chapter).

data generated by other lipidomic techniques.

reflects such variables for years [28].

**2.1 Gas chromatography: application in biological assays**

into their respective free fatty esters with lower boiling points (transesterification/ esterification reaction) [23]. Several methods are available for FAs derivatization [24], and the most applied ones are described in the 969.33 AOAC's method [25]. Particularly for cholesterol analysis, the samples preparation must consider a derivatization reaction. This allows to block protic sites of steroids obtained after an unsaponifiable lipid extraction [26] had been performed, and also, to diminish dipole-dipole interactions, to increase the volatility of the compounds, and to generate products with reduced polarity. Cholesterol derivatization is usually achieved by using trimethylsilyl compounds as reagents (silylation reaction). A common method for this purpose is described by Bowden and collaborators [27], in which *N*,*O*-bis(trimethylsilyl-trifluoroacetamide/trimethylchlorosilane)—BSTFA/

Nowadays, other more modern analytical tools than GC (next-generation techniques) do not require sample derivatization for lipid analysis. Needed lipid derivatization can be then consider a quite limitation step of the technique. In comparison with next-generation techniques, GC also implies in using greater sample quantities. This may be the main limitation in biological assays, which usually lead with restricted sample amounts. Nevertheless, by using certified standard and a powerful detector as FID, GC has the advantage to allow a precise and complete (by burning every compound, no one is lost in the detection) quantification of lipid compounds from biological samples, not always achieved by the other analytical techniques. In this context, GC continues to be accepted as an efficient and simple technique for FA and sterol analyses, mainly when combined with mass spectrom-

In biological issues, GC is largely applied to assess the FA and cholesterol contents in animal models or human fluids and tissues, as biological markers of FA ingestion and cell incorporation. The technique is a powerful tool in studies assessing the effect of FA supplementation on a specific biologic response. For instance, the endogenous synthesis of EPA and DHA from ALA is low in humans, who have in the ingestion, oily fishes as the most relevant source. Therefore, studies on n-3 PUFAs have been focused on the effect of fish ingestion or fish oil/EPA/DHA supplementation in several clinical conditions, and cell and disease models. In such studies, the treatment compliance or effectiveness can be reflected by the cell or circulate contends of n-3 PUFAs [28]. Furthermore, GC can be applied to validate

A practical example in using GC for treatment compliance is the study of Nogueira et al. [29] assessing the effect of n-3 PUFAs supplementation in patients with nonalcoholic steatohepatitis against placebo (mineral oil). In this study, GC highlighted a similar increase in n-3 PUFAs plasma in both n-3 PUFAs- and placebo-treated patients. Because the authors have controlled compliance of n-3 PUFAs, they were able to discover off-protocol intake of PUFAs by some patients from the placebo group. When studying biochemical markers of lipid intake and cell incorporation, the biological sample nature matters. For instance, plasma and red blood samples can reflect periods of weeks and months of FAs ingestion and their effects, respectively, while the adipose tissue is the reference method, once it

The study of Ravacci et al. [30] can illustrate the use of GC to assess treatment compliance. Applying this technique, the authors were able to demonstrate that the treatment of a lineage of breast cancer cell overexpressing HER-2 with DHA increased its availability in the cell membrane and was associated with the

**26**

disruption of surface lipid rafts that sustained cell signal for survival. Regarding the use of GC for data validation, a practical example is the study of Ouldamer et al. [31], which applied the technique to validate the fatty acid information generated by the 1 H NMR analysis on the PUFAs n-3 DHA and EPA content in the adipose tissue of mammary tumor model in rats exposed to controlled dietary intake of lipids.

## **3. Mass spectrometry: principles, strengths, and weaknesses**

Modernization of MS used for lipid analysis raised the concept of lipidomics. Lipidomics is an emerging science that aims to analyze the total lipid content found in a cell or tissue (lipidome) through the application of analytical chemistry principles and techniques. As a part of the omics sciences, the processes applied in the lipidomic analysis are analogous to those applied in other life-building macromolecules, such as deoxyribonucleic acid (DNA), ribonucleic acid (RNA), and proteins, called as genomics, transcriptomics, and proteomics, respectively [32].

The basic principle of the MS technique is founded on the detection of the abundance of ions by their mass/charge ratio (m/z). To allow the analysis, such ions of compounds are generated by suitable methods and ions are separated according to their m/z. Ionization techniques can break some sample's molecules into charged fragments and are chosen according to the physical state of the sample. Also, the efficiency of various ionization mechanisms for the unknown species might help when picking the most appropriate ionization technique. The most common ones for gaseous samples are the electron and chemical ionizations, while for liquid and solid samples, the electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) are usually applied [33].

Advances in ESI-MS and MALDI-MS have greatly facilitated lipidomic analysis [34] and enabled a great progress in lipid metabolic discoveries. This is because ESI is one of the softest ionization techniques, in which some complex dimers and solvent adducts can also be detected at the end. The efficiency of lipid ionization in ESI is incomparably higher than achieved by other traditional MS ion sources. MALDI-MS counts on a good solubility of analytes (lipids) and a matrix (for example, 2,5-dihydrobenzoic acid) in organic solvents, and provides excellent signal-to-noise ratio and reproducibility [32].

Mass spectrometers are made from three components: the ion source (1), which converts a sample into ions that are targeted through the mass analyzer (2) and run into the detector (3). The mass analyzer acts as ions organizer (classifier) using ion m/z ratios. This component accelerates ions as they face a strong electromagnetic field. The detector measures charged particles, such as an electron multiplier [35], and the abundances of each ion present in a sample are reported.

An advantage of MS is its high sensibility. A detection limit, expressed in concentration units, goes from a mol L<sup>−</sup><sup>1</sup> to as low as fmol L<sup>−</sup><sup>1</sup> and surely shall improve as the instruments modernize. For example, the instrument response factor for any individual molecular species detected is essentially identical within experimental error after 13C deisotoping if the analysis is performed properly [34]. Also, MS ion source can act as a separation device if set to selectively ionize just a certain lipid class. Thus, it is feasible to analyze different classes of lipids and individual molecular species with high efficiency without prior chromatographic separation. Nevertheless, depending on the analysis aims, MS can also be combined with a chromatography system, as GC (early mentioned) and liquid chromatography (LC) [35].

Data obtained by MS are displayed as spectra of the relative abundance of detected ions as a function of the corresponding m/z. By correlating the known masses (e.g., an entire molecule) to the identified masses, or through the compounds deposited characteristic fragmentation pattern, MS are used to identify compounds. The MS are also used to determine the elemental or isotopic signature of a sample, the masses of particles and molecules, and to elucidate the chemical structures of molecules [36]. Database platforms, such as LIPIDMAPS, LIPID Bank, LIPIDAT, Cyberlipids, and Lipidomics expertise platform, can help to identify the lipid molecules. Then, interpretation of MS-obtained lipid data must be conducted in accordance with the literature [7].

When assessing the entire lipidome profile, i.e., lipidomics by MS or nuclear magnetic resonance (NMR, detailed later in this chapter), big-data information is generated. Therefore, lipidomics require multistatistic tools for data interpretation. The additional information to MS lipidomics is mapping of the lipid pathway. For example, diacylglycerol is an essential precursor for glycerophospholipid and glycerolipid synthesis in eukaryotes [5].

Manual data interpretation using publicly available databases (i.e., KEGG pathways and the LipidMAPS databases) may add in to lipidomic results and provide meaningful biological context to data understanding from biological point of view. Indeed, using bioinformatics software platform, one can understand the changes in lipid composition and content, and understand adaptive or pathological changes in lipid metabolism. Lipids form networks, which are used to build their inter-relationships and connect them based on known metabolic pathways. Also, these relationships and the determined quantities of lipids are used to calculate the possible contributions to the production of a particular lipid class in the network, and the masses calculated are compared with the masses determined from the lipidomic MS data.

Several parameters involving the metabolic pathways can then be derived from computational simulation, such as those associated with enzymatic activities, as those analyzed by a lipid expertise, i.e., known principles of lipid biochemistry to calculate indexes of fatty acid unsaturation, fatty acyl chain length, or fatty acid precursor/product ratios to gain insight into the function of fatty acid remodeling or other relevant lipid metabolic pathways [5, 37]. Some useful tools that can be used for this purpose are the public platforms MetaboAnalyst (available from http://www.metaboanalyst.ca), VANTED, and MAVEN [37].

Once lipids have a high discrepancy of m/z within their categories and are susceptible to ion cleavage, the main disadvantage of MS in lipid analysis is that some compounds from a mixture may be determined as the same ion and incorrectly identified. Furthermore, lipid quantification by MS may be weakened by the loss of ion information due to the random collision of lipid molecules that may preclude that all of these get to the detector, the differing abilities of lipid species to form ions and hence varying signal intensity, and the ion-quenching phenomena. The last can occur when the signal from poor ionizing lipids is quenched by more easily ionized species (therefore suppressing the former signal), which is quite avoided by prior separation of lipid species for accurate quantitation or the use of specialized MS [38]. Altogether, these factors result in a loss of sensitivity for some nonpolar lipid metabolites.

It is worth to note that the limitations in identification and quantification of lipid species by MS described above have been minimized with advances of the technique (i.e., target MS). Currently, this analytic tool is considered accurate for characterization of lipids and the most efficient one to assess lipidomes.

#### **3.1 Mass spectrometry: application in biological assays**

In biological assays, lipidomics-MS analysis is highly applied to generate information related to metabolism and biological responses, once several known

**29**

performed.

**and weaknesses**

*Analytical Tools for Lipid Assessment in Biological Assays*

diseases (in comparison with healthy status) [5, 32, 39].

pathways from metabolic networks in eukaryotes involve lipids as metabolic intermediates (mainly sphingolipids, glycerophospholipids, glycerolipids, and nonesterified fatty acids [NEFAs]) or signaling molecules (mainly oxysterols) [5]. For instance, changes of a lipidome profile can be identified by MS, allowing the interpretation of biological responses to external interferences (i.e., by comparing the lipidome before and after a medication) or enrolled in the pathophysiology of

The ionization technique applied is a relevant point to be considered when designing studies for lipid assessment in biological samples using MS. For instance, MALDI can be used to analyze changes of lipid and their metabolites in single genetically identical cells from the RAW264.7 lineage after lipopolysaccharide (LPS) stimulation, using a Fourier transform ion cyclotron resonance mass spectrometer (FTICR MS). MALDI analysis was chosen because single cells on a plate using a histology-directed workflow can increase the number of cells analyzed. Furthermore, the speed of MALDI-IMS enables high spatial resolution and highthroughput single-cell analysis. Combined with the high sensitivity of FTICR MS, hundreds of lipids can be measured from a large population of single cells (>100) in

Tandem MS measurements (i.e., through precursor ion scanning and neutral loss scanning experiments) are useful for biological assays requiring the identification of all lipid molecular species. These methodologies are usually better than full scan MS because they apply sequential analyzers and are often associated with a target analysis (i.e., aiming to study a molecule species). This allows high sensitivity and enhanced signal/noise ratio, facilitating the characterization of minor but

One example of the tandem analysis application is the work of Slatter et al. [41]. By using LC-MS/MS (tandem MS), they were able to characterize the lipidomic network of human platelets, where nearly 200 oxidized species were identified. These minacious data provided by the methodology allowed to display a direct link between innate immunity and mitochondrial bioenergetics in human platelets. Procedures enabling to achieve this conclusion from generated data included the selection of lipids upregulated under thrombin activation and the analysis on temporal dynamics of their generation, monitoring precursor-to-product ion

Also, through tandem MS, Morgan et al. [42] have proposed a novel role for 12/15-lipoxygenase in regulating autophagy. They have used LC/ESI/MS/ MS in a target approach to determine the levels of 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (DMPE), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dimyristoyl-sn-glycero-3-phosphate (DMPA), and 1,2-dimyristoylsn-glycero-3-phospho-(1′-rac-glycerol) (DMPG), using comparison technique with internal standards. In addition, the 1,2-dimyristoyl-sn-glycero-3-phosphoserine (DMPS) was determined by product ions and the analysis of cholesteryl esters was

**4. Nuclear magnetic resonance spectroscopy: principles, strengths,** 

Along with other analytical tools available for lipidome investigations, NMR spectroscopy allows identification of characteristic signals from the different classes of lipids and provides their successful quantification [43, 44]. The technique facilitates the analysis of hundreds of metabolites in a single sample with great advantage

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

biologically relevant lipid species [40].

transitions in multiple reaction monitoring (MRM) modes.

because there is no need for a previous sample treatment [8].

a few hours.

#### *Analytical Tools for Lipid Assessment in Biological Assays DOI: http://dx.doi.org/10.5772/intechopen.81523*

*Advances in Lipid Metabolism*

in accordance with the literature [7].

glycerolipid synthesis in eukaryotes [5].

lipidomic MS data.

lipid metabolites.

compounds deposited characteristic fragmentation pattern, MS are used to identify compounds. The MS are also used to determine the elemental or isotopic signature of a sample, the masses of particles and molecules, and to elucidate the chemical structures of molecules [36]. Database platforms, such as LIPIDMAPS, LIPID Bank, LIPIDAT, Cyberlipids, and Lipidomics expertise platform, can help to identify the lipid molecules. Then, interpretation of MS-obtained lipid data must be conducted

When assessing the entire lipidome profile, i.e., lipidomics by MS or nuclear magnetic resonance (NMR, detailed later in this chapter), big-data information is generated. Therefore, lipidomics require multistatistic tools for data interpretation. The additional information to MS lipidomics is mapping of the lipid pathway. For example, diacylglycerol is an essential precursor for glycerophospholipid and

Manual data interpretation using publicly available databases (i.e., KEGG pathways and the LipidMAPS databases) may add in to lipidomic results and provide meaningful biological context to data understanding from biological point of view. Indeed, using bioinformatics software platform, one can understand the changes in lipid composition and content, and understand adaptive or pathological changes in lipid metabolism. Lipids form networks, which are used to build their inter-relationships and connect them based on known metabolic pathways. Also, these relationships and the determined quantities of lipids are used to calculate the possible contributions to the production of a particular lipid class in the network, and the masses calculated are compared with the masses determined from the

Several parameters involving the metabolic pathways can then be derived from computational simulation, such as those associated with enzymatic activities, as those analyzed by a lipid expertise, i.e., known principles of lipid biochemistry to calculate indexes of fatty acid unsaturation, fatty acyl chain length, or fatty acid precursor/product ratios to gain insight into the function of fatty acid remodeling or other relevant lipid metabolic pathways [5, 37]. Some useful tools that can be used for this purpose are the public platforms MetaboAnalyst (available from

Once lipids have a high discrepancy of m/z within their categories and are susceptible to ion cleavage, the main disadvantage of MS in lipid analysis is that some compounds from a mixture may be determined as the same ion and incorrectly identified. Furthermore, lipid quantification by MS may be weakened by the loss of ion information due to the random collision of lipid molecules that may preclude that all of these get to the detector, the differing abilities of lipid species to form ions and hence varying signal intensity, and the ion-quenching phenomena. The last can occur when the signal from poor ionizing lipids is quenched by more easily ionized species (therefore suppressing the former signal), which is quite avoided by prior separation of lipid species for accurate quantitation or the use of specialized MS [38]. Altogether, these factors result in a loss of sensitivity for some nonpolar

It is worth to note that the limitations in identification and quantification of lipid species by MS described above have been minimized with advances of the technique (i.e., target MS). Currently, this analytic tool is considered accurate for

In biological assays, lipidomics-MS analysis is highly applied to generate information related to metabolism and biological responses, once several known

characterization of lipids and the most efficient one to assess lipidomes.

**3.1 Mass spectrometry: application in biological assays**

http://www.metaboanalyst.ca), VANTED, and MAVEN [37].

**28**

pathways from metabolic networks in eukaryotes involve lipids as metabolic intermediates (mainly sphingolipids, glycerophospholipids, glycerolipids, and nonesterified fatty acids [NEFAs]) or signaling molecules (mainly oxysterols) [5]. For instance, changes of a lipidome profile can be identified by MS, allowing the interpretation of biological responses to external interferences (i.e., by comparing the lipidome before and after a medication) or enrolled in the pathophysiology of diseases (in comparison with healthy status) [5, 32, 39].

The ionization technique applied is a relevant point to be considered when designing studies for lipid assessment in biological samples using MS. For instance, MALDI can be used to analyze changes of lipid and their metabolites in single genetically identical cells from the RAW264.7 lineage after lipopolysaccharide (LPS) stimulation, using a Fourier transform ion cyclotron resonance mass spectrometer (FTICR MS). MALDI analysis was chosen because single cells on a plate using a histology-directed workflow can increase the number of cells analyzed. Furthermore, the speed of MALDI-IMS enables high spatial resolution and highthroughput single-cell analysis. Combined with the high sensitivity of FTICR MS, hundreds of lipids can be measured from a large population of single cells (>100) in a few hours.

Tandem MS measurements (i.e., through precursor ion scanning and neutral loss scanning experiments) are useful for biological assays requiring the identification of all lipid molecular species. These methodologies are usually better than full scan MS because they apply sequential analyzers and are often associated with a target analysis (i.e., aiming to study a molecule species). This allows high sensitivity and enhanced signal/noise ratio, facilitating the characterization of minor but biologically relevant lipid species [40].

One example of the tandem analysis application is the work of Slatter et al. [41]. By using LC-MS/MS (tandem MS), they were able to characterize the lipidomic network of human platelets, where nearly 200 oxidized species were identified. These minacious data provided by the methodology allowed to display a direct link between innate immunity and mitochondrial bioenergetics in human platelets. Procedures enabling to achieve this conclusion from generated data included the selection of lipids upregulated under thrombin activation and the analysis on temporal dynamics of their generation, monitoring precursor-to-product ion transitions in multiple reaction monitoring (MRM) modes.

Also, through tandem MS, Morgan et al. [42] have proposed a novel role for 12/15-lipoxygenase in regulating autophagy. They have used LC/ESI/MS/ MS in a target approach to determine the levels of 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (DMPE), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dimyristoyl-sn-glycero-3-phosphate (DMPA), and 1,2-dimyristoylsn-glycero-3-phospho-(1′-rac-glycerol) (DMPG), using comparison technique with internal standards. In addition, the 1,2-dimyristoyl-sn-glycero-3-phosphoserine (DMPS) was determined by product ions and the analysis of cholesteryl esters was performed.

## **4. Nuclear magnetic resonance spectroscopy: principles, strengths, and weaknesses**

Along with other analytical tools available for lipidome investigations, NMR spectroscopy allows identification of characteristic signals from the different classes of lipids and provides their successful quantification [43, 44]. The technique facilitates the analysis of hundreds of metabolites in a single sample with great advantage because there is no need for a previous sample treatment [8].

The principle of NMR spectroscopy is based on the physical resonance phenomenon in which spin-active nuclei in a strong static magnetic field respond to a perturbation (radiofrequency waves) by producing an electromagnetic signal with a characteristic frequency, which matches magnetic field observed by a given nucleus. This process of resonance happens when the oscillation frequency matches the intrinsic frequency of the nuclei, which depends on the strength of the static magnetic field, the chemical environment, and the magnetic properties of the isotope involved [45].

In a practical way, NMR spectroscopy provides information of the number (integrals) of magnetically distinct atoms (chemical shift of the resonance frequencies and peak splitting due to the coupling constants J or dipolar couplings between nuclear spins in the sample) of the studied isotope and provides all necessary information for determination of the structure of unknown molecules. Several nuclei can be studied by NMR techniques, but the most commonly available ones are hydrogen-1 and carbon-13. The most common experiments for lipid analysis by NMR are 1 H, 13C, 31P, and the bidimensional experiments involving 1 H-1 H and 1 H-13C [45].

Usually, an NMR experiment starts with insertion of a liquid sample into the magnet, then, short radio-frequency pulses (from an electronic device named probe) are applied, and all emitted frequencies from the same type of nuclei are registered and reported as signals with a given chemical shift, multiplicity, and intensity. Also, multidimensional NMR as well as solid-state NMR has emerged to provide additional and relevant information on sample composition [45].

Also, the exact ratio of specific fatty acids in the lipid samples and their iodine values could be calculated considering integral values corresponding to characteristic peaks with the help of the corresponding spectral information and the existing references [46]. This type of experiment works as a relative quantification. Absolute and relative quantification experiments by NMR are possible; however, it is necessary to take care of some precautions. Direct quantitative information by NMR is due to the fact that the signal intensity of each resonance in the NMR spectrum is directly proportional to the number of spins associated with the particular resonance [38]. Thus, no standard with chemical similarity to the studied compounds is required as in other analytical methods; however, one certified standard must be used. This can be performed through relative quantification using ERETIC. For absolute quantification also, a certified standard is required now as an internal standard in a known concentration. For both methods, the pulse sequence needs to be calibrated to 90° to be sure that the spectral response is completely real, and it means that the longitudinal relaxation time (T1) of spins is entirely returned [38]. Typically, this is achieved by waiting five times the longest T1 (at five times T1 approximately 99.3% of the equilibrium value is re-established) between two scans.

Proton magnetic resonance spectroscopic imaging (1 H-MRSI) has a major role in lipid assays, mainly used in the medical area with extreme importance for *in vivo* sampling. Both profiling and ratio quantifications are possible by the obtained spatial resolved spectra. The presence of so many compounds in living biological samples may require water or other signal suppression experiments to be performed in order to obtain better resolution on the target metabolites. The same approach is used in NMR samples but with greater implications due to lack of sample pretreatment [47].

Compared to the MS method, NMR technique is less sensitive and limited by the overlapping of signals in either, <sup>1</sup> H NMR or 31P NMR, and also by the low natural abundance of 13C for 13C NMR. On the other hand, NMR is a nondestructive sample technique that allows a high analytical reproducibility, an easy identification of molecular moieties, and with relatively easy to get information on molecular dynamics [8, 38]. Furthermore, NMR does not require a standard curve or molecule species for quantitative measuring. Therefore, this technique has been emerging as

**31**

*Analytical Tools for Lipid Assessment in Biological Assays*

a promising approach for more accurate and faster quantitative analysis of lipids than other analytical methods [38]. Also, the sensitivity improvement of cryogenic probe in an equipment of 800 MHz LC-NMR is very promising in analysis of a trace

of approximately 1 μg sample within 30 min, whereas the current 500 MHz NMR

**4.1 Nuclear magnetic resonance spectroscopy: application in biological assays**

issues where biofluid samples such as serum, plasma, urine, cerebrospinal fluid

experiments, which bring rich information on lipid profiling, for example, molecular identification of fatty acid chains and phospholipid structures. Furthermore, heteronuclear and multidimensional experiments can be used to elucidate lipid profiling information by signal interpretation and also using comparisons with databases. The 13C NMR is also a complementary tool that can be used for fatty acyl

Once NMR allows the noninvasive lipid analysis in intact cells and tissues, the technique prevents losses of chemical information in the analyte environment. This fact, together with the high sensibility of NMR to molecular dynamics (in timescales from picoseconds to seconds), enables to investigate changes in lipid and dynamic structures in biochemical cell functionalization. The experiment used for this application is the diffusion ordered spectroscopy (DOSY), which enables to separate signals according to their diffusion coefficients and then add chromatography-

Lipoproteins consist mainly from cholesterol esters and triacylglycerols surrounded by a hydrophilic layer, which comprehend phospholipids, cholesterol, and proteins [8]. Lipoproteins perform the lipid transportation in blood circulation through the exogenous (dietary lipids) and the endogenous (liver-synthetized lipids) channels. The endogenous transportation begins in the liver through the production of a very low-density lipoprotein (VLDL). After being secreted into the bloodstream, VLDL interacts with other lipoproteins, through collisions, in which

Kostara et al. [49] have found how blood lipoproteins influence the progression of coronary heart disease (CHD) by comparing the lipid profiles of atherogenic (non-HDL) and atheroprotective (HDL) lipoproteins from patients with CHD with those from patients with normal coronary arteries (NCA). They analyzed the lipid extracts

fied the potential target-lipid biomarkers for the early evaluation of the CHD onset. Furthermore, Lopes et al. [50] were able to find that circulating HDL increases, and LDL and VLDL decrease in obese patients after bariatric surgery by using DOSY experiments to monitor these lipoproteins. Notably, lipoprotein investigations and quantitative analysis of lipids can be performed using NMR of the same sample [51]. Also, selective recoupling of dipolar and chemical-shift interactions removed by magic-angle spinning NMR in the solid state allows the characterization of regulatory interactions, dynamics, and ion channels within biological membranes [52]. In this scenario, the NMR application has contributed to obtaining of important data on the structure and turnover of lipid species and the composition of lipids in cells, and to characterize pathways enrolled in lipid synthesis/transport and degradation [53, 54]. Also, the high-resolution magic-angle spinning NMR (HR-MAS

H NMR experiments and statistical analysis and identi-

the contact with the high-density lipoprotein (HDL) is highlighted.

NMR) has been applied to global lipidomic studies [52].

A wide variety of NMR experiments (e.g., HSQC, HMBC, TOCSY, etc.) besides

H, 13C, and 31P NMR are being used to solve a variety of biological

H NMR spectrum

H, 13C, and 31P NMR

amount of lipids in a faster experiment, once it is able to acquire <sup>1</sup>

(CSF), etc., are being investigated. More commonly used are 1

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

needs 20 h or longer [38].

residue identification [38].

like capabilities to NMR [38, 48].

of these lipoproteins using 1

the classics <sup>1</sup>

*Advances in Lipid Metabolism*

isotope involved [45].

1

The principle of NMR spectroscopy is based on the physical resonance phenomenon in which spin-active nuclei in a strong static magnetic field respond to a perturbation (radiofrequency waves) by producing an electromagnetic signal with a characteristic frequency, which matches magnetic field observed by a given nucleus. This process of resonance happens when the oscillation frequency matches the intrinsic frequency of the nuclei, which depends on the strength of the static magnetic field, the chemical environment, and the magnetic properties of the

In a practical way, NMR spectroscopy provides information of the number (integrals) of magnetically distinct atoms (chemical shift of the resonance frequencies and peak splitting due to the coupling constants J or dipolar couplings between nuclear spins in the sample) of the studied isotope and provides all necessary information for determination of the structure of unknown molecules. Several nuclei can be studied by NMR techniques, but the most commonly available ones are hydrogen-1 and carbon-13. The most common experiments for lipid analysis by NMR are

Usually, an NMR experiment starts with insertion of a liquid sample into the magnet, then, short radio-frequency pulses (from an electronic device named probe) are applied, and all emitted frequencies from the same type of nuclei are registered and reported as signals with a given chemical shift, multiplicity, and intensity. Also, multidimensional NMR as well as solid-state NMR has emerged to

Also, the exact ratio of specific fatty acids in the lipid samples and their iodine values could be calculated considering integral values corresponding to characteristic peaks with the help of the corresponding spectral information and the existing references [46]. This type of experiment works as a relative quantification. Absolute and relative quantification experiments by NMR are possible; however, it is necessary to take care of some precautions. Direct quantitative information by NMR is due to the fact that the signal intensity of each resonance in the NMR spectrum is directly proportional to the number of spins associated with the particular resonance [38]. Thus, no standard with chemical similarity to the studied compounds is required as in other analytical methods; however, one certified standard must be used. This can be performed through relative quantification using ERETIC. For absolute quantification also, a certified standard is required now as an internal standard in a known concentration. For both methods, the pulse sequence needs to be calibrated to 90° to be sure that the spectral response is completely real, and it means that the longitudinal relaxation time (T1) of spins is entirely returned [38]. Typically, this is achieved by waiting five times the longest T1 (at five times T1 approximately 99.3% of the equilibrium value is re-established) between two scans.

in lipid assays, mainly used in the medical area with extreme importance for *in vivo* sampling. Both profiling and ratio quantifications are possible by the obtained spatial resolved spectra. The presence of so many compounds in living biological samples may require water or other signal suppression experiments to be performed in order to obtain better resolution on the target metabolites. The same approach is used in NMR samples but with greater implications due to lack of sample pretreatment [47]. Compared to the MS method, NMR technique is less sensitive and limited

natural abundance of 13C for 13C NMR. On the other hand, NMR is a nondestructive sample technique that allows a high analytical reproducibility, an easy identification of molecular moieties, and with relatively easy to get information on molecular dynamics [8, 38]. Furthermore, NMR does not require a standard curve or molecule species for quantitative measuring. Therefore, this technique has been emerging as

provide additional and relevant information on sample composition [45].

H-1

H and 1

H-MRSI) has a major role

H NMR or 31P NMR, and also by the low

H-13C [45].

H, 13C, 31P, and the bidimensional experiments involving 1

Proton magnetic resonance spectroscopic imaging (1

by the overlapping of signals in either, <sup>1</sup>

**30**

a promising approach for more accurate and faster quantitative analysis of lipids than other analytical methods [38]. Also, the sensitivity improvement of cryogenic probe in an equipment of 800 MHz LC-NMR is very promising in analysis of a trace amount of lipids in a faster experiment, once it is able to acquire <sup>1</sup> H NMR spectrum of approximately 1 μg sample within 30 min, whereas the current 500 MHz NMR needs 20 h or longer [38].

## **4.1 Nuclear magnetic resonance spectroscopy: application in biological assays**

A wide variety of NMR experiments (e.g., HSQC, HMBC, TOCSY, etc.) besides the classics <sup>1</sup> H, 13C, and 31P NMR are being used to solve a variety of biological issues where biofluid samples such as serum, plasma, urine, cerebrospinal fluid (CSF), etc., are being investigated. More commonly used are 1 H, 13C, and 31P NMR experiments, which bring rich information on lipid profiling, for example, molecular identification of fatty acid chains and phospholipid structures. Furthermore, heteronuclear and multidimensional experiments can be used to elucidate lipid profiling information by signal interpretation and also using comparisons with databases. The 13C NMR is also a complementary tool that can be used for fatty acyl residue identification [38].

Once NMR allows the noninvasive lipid analysis in intact cells and tissues, the technique prevents losses of chemical information in the analyte environment. This fact, together with the high sensibility of NMR to molecular dynamics (in timescales from picoseconds to seconds), enables to investigate changes in lipid and dynamic structures in biochemical cell functionalization. The experiment used for this application is the diffusion ordered spectroscopy (DOSY), which enables to separate signals according to their diffusion coefficients and then add chromatographylike capabilities to NMR [38, 48].

Lipoproteins consist mainly from cholesterol esters and triacylglycerols surrounded by a hydrophilic layer, which comprehend phospholipids, cholesterol, and proteins [8]. Lipoproteins perform the lipid transportation in blood circulation through the exogenous (dietary lipids) and the endogenous (liver-synthetized lipids) channels. The endogenous transportation begins in the liver through the production of a very low-density lipoprotein (VLDL). After being secreted into the bloodstream, VLDL interacts with other lipoproteins, through collisions, in which the contact with the high-density lipoprotein (HDL) is highlighted.

Kostara et al. [49] have found how blood lipoproteins influence the progression of coronary heart disease (CHD) by comparing the lipid profiles of atherogenic (non-HDL) and atheroprotective (HDL) lipoproteins from patients with CHD with those from patients with normal coronary arteries (NCA). They analyzed the lipid extracts of these lipoproteins using 1 H NMR experiments and statistical analysis and identified the potential target-lipid biomarkers for the early evaluation of the CHD onset. Furthermore, Lopes et al. [50] were able to find that circulating HDL increases, and LDL and VLDL decrease in obese patients after bariatric surgery by using DOSY experiments to monitor these lipoproteins. Notably, lipoprotein investigations and quantitative analysis of lipids can be performed using NMR of the same sample [51].

Also, selective recoupling of dipolar and chemical-shift interactions removed by magic-angle spinning NMR in the solid state allows the characterization of regulatory interactions, dynamics, and ion channels within biological membranes [52].

In this scenario, the NMR application has contributed to obtaining of important data on the structure and turnover of lipid species and the composition of lipids in cells, and to characterize pathways enrolled in lipid synthesis/transport and degradation [53, 54]. Also, the high-resolution magic-angle spinning NMR (HR-MAS NMR) has been applied to global lipidomic studies [52].

Besides the identification of lipid species and dynamics, NMR can be used for reliable quantification of lipid mixtures obtained from tissues, body fluids, and cell cultures [40, 55]. It can be allied to the bioinformatic tools available to a better quantitative analysis of lipid profiles [56]. For instance, using 1 H NMR and 31P NMR, Fernando et al. [57] were able to identify an over-accumulation of lipids associated with the pathophysiology of ethanol-induced liver steatosis accompanied by mild inflammation.

Also, quantification can be used in magnetic resonance imaging (MRI) experiments as Vafaeyan et al. [47] have shown. They have used a time-domain quantification method namely as subtract-QUEST-MRSI algorithm to quantify alterations of the biomarkers, i.e., lipids and other metabolism molecules species such as choline, creatine, *N*-acetyl aspartate, lactate, myo-inositol, and glutamine in multiple sclerosis subjects in comparison with control group. This research aimed to know how lesion biomarker ratios in multiple sclerosis have affected human brains, through the imaging of different brain areas, which could present lesions.

Other MRI works have found that on brain imaging, lipids tend to be an almost undesired artifact, and consequently, scientists may use the approach of selective signal suppression pulses such as adiabatic frequency selective, spatial-spectral lipid suppression, or broadband outer volume suppression bands [58]. Trauner et al. [59] have used a dynamic saturation transfer technique in MRI experiments to assess dynamic Pi-to-ATP exchange parameters in nonalcoholic fatty liver disease (NAFLD) and steatohepatitis (NASH) aiming to report alterations of hepatic lipid, cell membrane, and energy.

## **5. Final considerations**

Lipids *per se* exert several relevant biological functions making the single knowledge of the lipidome profile from a biological sample highly informative by itself. For instance, sphingolipids and glycerophospholipids are important components of the cell membrane and then can affect several cellular functions. Disorders of sphingolipid metabolism are associated with lysosomal storage diseases and of lysoglycerophospholipid by phospholipase A2 activation are associated with lipotoxicity and inflammation. Accumulation of triacylglycerol (a glycerophospholipid) is associated with lipotoxicity and insulin resistance, and the NEFA profile is a useful indicator of lipid metabolism and can add to understanding on molecular mechanisms underlying the metabolic syndrome [5].

Therefore, lipidomic tools are particularly useful to identify and understand changes in metabolic pathways and the underlying mechanisms enrolled in the pathophysiology of human health, such as metabolic diseases. One practical example is data from Meikle et al. [60] study that measured 259 lipid species in plasma samples from prediabetic, diabetic and normal glucose tolerant patients, including sphingolipids, phospholipids, glycerolipids, and cholesterol ester. The authors used electrospray ionization-tandem mass spectrometry in previous precursor ion and neutral loss scans on control plasma extracts, MRM experiments for the major species of each lipid class identified in plasma, and quantification using internal standards. These approaches highlighted that metabolic pathways altered in type 2 diabetes include a deregulation of lipid homeostasis, characterized by abnormal plasma-free fatty acids accumulation.

In lipidomic studies, beyond the care of equipment calibration and accuracy of the experiments, special cares of analytical procedures must be planned to have accurate information of data. The statistical recourses are necessary to process the data, but, also lipid knowledge is required for correct interpretation in all cases. The

**33**

*Analytical Tools for Lipid Assessment in Biological Assays*

choice of the most suitable lipidomic tool to be used for a specific biological assay is closely linked to the study aim. Next-generation techniques (MS and NMR) can provide detailed lipid information to assess more elaborated scientific questions. However, thousands of individual lipid molecular species are present in cells implying that no single technique can effectively study all the lipid species [38]. When possible, combining techniques can be the best choice, because one can compensate for the limitation of the other, and bring complementary information and/or can

Usually, combined lipidomic techniques are applied for data validation. For instance, data obtained by shotgun lipidomics (direct infusion MS) can be validated using LC-MS-based analyses and *vice versa*. Other methods, including NMR, or chromatography-based analysis might be used to validate the total lipid content of a lipid class [5]. However, the combined use of lipidomic techniques can also be useful to improve the data information on the lipids from biological samples. For instance, to assess lipid changes during the response of hypoxia stress to a treatment in cervical cancer-derived cells (HeLa cells), Yu et al. [40] applied NMR technique for the phospholipid profile analysis and MS for phospholipids characterization. Also, Whiley et al. [61] investigated the plasma phosphatidylcholine metabolism using NMR and MS to obtain a fingerprint of three phosphatidylcholines (PC) molecules that significantly decrease in individuals with Alzheimer's disease compared to healthy controls. Then, LC-MS and NMR were used for phosphatidylcholine and fatty acyl side chain validation and for total plasma choline validation, respectively. The study of Whiley et al. [61] illustrates the scientific value in combining different lipidomic tools to obtain complementary information and reinforce validation of

In conclusion, all available tools for lipidomic studies in biological samples have several advantages and limitations that can be overcome when combining more than one technique. Because this practice involves the availability of complex technologies and skilled labor, it is not always possible. In this scenario, the use of mass spectrometry alone can be the best alternative currently available when technique combination is impossible. However, NMR has a high potential and, in the future,

This research counted on grant received from the Brazilian agency—*Fundação* 

*de Amparo à Pesquisa do Estado de São Paulo*—FAPESP (Sao Paulo Research

may be expected to answer issues that MS is quite limited to do.

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

validate the previous analysis data.

the obtained data.

**Acknowledgements**

**Conflict of interest**

Foundation, grant number 2018/06510-4).

Authors declare no conflict of interests.

#### *Analytical Tools for Lipid Assessment in Biological Assays DOI: http://dx.doi.org/10.5772/intechopen.81523*

*Advances in Lipid Metabolism*

by mild inflammation.

cell membrane, and energy.

**5. Final considerations**

Besides the identification of lipid species and dynamics, NMR can be used for reliable quantification of lipid mixtures obtained from tissues, body fluids, and cell cultures [40, 55]. It can be allied to the bioinformatic tools available to a bet-

NMR, Fernando et al. [57] were able to identify an over-accumulation of lipids associated with the pathophysiology of ethanol-induced liver steatosis accompanied

Also, quantification can be used in magnetic resonance imaging (MRI) experiments as Vafaeyan et al. [47] have shown. They have used a time-domain quantification method namely as subtract-QUEST-MRSI algorithm to quantify alterations of the biomarkers, i.e., lipids and other metabolism molecules species such as choline, creatine, *N*-acetyl aspartate, lactate, myo-inositol, and glutamine in multiple sclerosis subjects in comparison with control group. This research aimed to know how lesion biomarker ratios in multiple sclerosis have affected human brains,

through the imaging of different brain areas, which could present lesions.

Other MRI works have found that on brain imaging, lipids tend to be an almost undesired artifact, and consequently, scientists may use the approach of selective signal suppression pulses such as adiabatic frequency selective, spatial-spectral lipid suppression, or broadband outer volume suppression bands [58]. Trauner et al. [59] have used a dynamic saturation transfer technique in MRI experiments to assess dynamic Pi-to-ATP exchange parameters in nonalcoholic fatty liver disease (NAFLD) and steatohepatitis (NASH) aiming to report alterations of hepatic lipid,

Lipids *per se* exert several relevant biological functions making the single knowledge of the lipidome profile from a biological sample highly informative by itself. For instance, sphingolipids and glycerophospholipids are important components of the cell membrane and then can affect several cellular functions. Disorders of sphingolipid metabolism are associated with lysosomal storage diseases and of lysoglycerophospholipid by phospholipase A2 activation are associated with lipotoxicity and inflammation. Accumulation of triacylglycerol (a glycerophospholipid) is associated with lipotoxicity and insulin resistance, and the NEFA profile is a useful indicator of lipid metabolism and can add to understanding on molecular

Therefore, lipidomic tools are particularly useful to identify and understand changes in metabolic pathways and the underlying mechanisms enrolled in the pathophysiology of human health, such as metabolic diseases. One practical example is data from Meikle et al. [60] study that measured 259 lipid species in plasma samples from prediabetic, diabetic and normal glucose tolerant patients, including sphingolipids, phospholipids, glycerolipids, and cholesterol ester. The authors used electrospray ionization-tandem mass spectrometry in previous precursor ion and neutral loss scans on control plasma extracts, MRM experiments for the major species of each lipid class identified in plasma, and quantification using internal standards. These approaches highlighted that metabolic pathways altered in type 2 diabetes include a deregulation of lipid homeostasis, characterized by abnormal

In lipidomic studies, beyond the care of equipment calibration and accuracy of the experiments, special cares of analytical procedures must be planned to have accurate information of data. The statistical recourses are necessary to process the data, but, also lipid knowledge is required for correct interpretation in all cases. The

mechanisms underlying the metabolic syndrome [5].

plasma-free fatty acids accumulation.

H NMR and 31P

ter quantitative analysis of lipid profiles [56]. For instance, using 1

**32**

choice of the most suitable lipidomic tool to be used for a specific biological assay is closely linked to the study aim. Next-generation techniques (MS and NMR) can provide detailed lipid information to assess more elaborated scientific questions. However, thousands of individual lipid molecular species are present in cells implying that no single technique can effectively study all the lipid species [38]. When possible, combining techniques can be the best choice, because one can compensate for the limitation of the other, and bring complementary information and/or can validate the previous analysis data.

Usually, combined lipidomic techniques are applied for data validation. For instance, data obtained by shotgun lipidomics (direct infusion MS) can be validated using LC-MS-based analyses and *vice versa*. Other methods, including NMR, or chromatography-based analysis might be used to validate the total lipid content of a lipid class [5]. However, the combined use of lipidomic techniques can also be useful to improve the data information on the lipids from biological samples. For instance, to assess lipid changes during the response of hypoxia stress to a treatment in cervical cancer-derived cells (HeLa cells), Yu et al. [40] applied NMR technique for the phospholipid profile analysis and MS for phospholipids characterization. Also, Whiley et al. [61] investigated the plasma phosphatidylcholine metabolism using NMR and MS to obtain a fingerprint of three phosphatidylcholines (PC) molecules that significantly decrease in individuals with Alzheimer's disease compared to healthy controls. Then, LC-MS and NMR were used for phosphatidylcholine and fatty acyl side chain validation and for total plasma choline validation, respectively. The study of Whiley et al. [61] illustrates the scientific value in combining different lipidomic tools to obtain complementary information and reinforce validation of the obtained data.

In conclusion, all available tools for lipidomic studies in biological samples have several advantages and limitations that can be overcome when combining more than one technique. Because this practice involves the availability of complex technologies and skilled labor, it is not always possible. In this scenario, the use of mass spectrometry alone can be the best alternative currently available when technique combination is impossible. However, NMR has a high potential and, in the future, may be expected to answer issues that MS is quite limited to do.

## **Acknowledgements**

This research counted on grant received from the Brazilian agency—*Fundação de Amparo à Pesquisa do Estado de São Paulo*—FAPESP (Sao Paulo Research Foundation, grant number 2018/06510-4).

## **Conflict of interest**

Authors declare no conflict of interests.

*Advances in Lipid Metabolism*

## **Author details**

Banny Silva Barbosa Correia1 , Raquel Susana Torrinhas2 , William Yutaka Ohashi3,4 and Ljubica Tasic3 \*

1 Chemistry Institute, University of Sao Paulo (USP), Sao Carlos, Sao Paulo, Brazil

2 Department of Gastroenterology, Surgical Division (LIM 35), University of Sao Paulo (USP), School of Medicine, Sao Paulo, Brazil

3 Chemistry Institute, Campinas State University (UNICAMP), Campinas, Sao Paulo, Brazil

4 Agilent Technologies Brasil Ltda., Barueri, Sao Paulo, Brazil

\*Address all correspondence to: ljubica@iqm.unicamp.br

© 2018 The Author(s). Licensee IntechOpen. 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.

**35**

gene.2014.10.039

*Analytical Tools for Lipid Assessment in Biological Assays*

[9] Calder PC, Deckelbaum RJ. Dietary lipids: More than just a source of calories. Current Opinion in Clinical Nutrition and Metabolic Care.

[10] Calder PC. Marine omega-3 fatty acids and inflammatory processes: Effects, mechanisms and clinical relevance. Biochimica et Biophysica Acta. 2015;**1851**(4):469-484. DOI: 10.1016/j.bbalip.2014.08.010

[11] Muro E, Atilla-Gokcumen GE, Eggert US. Lipids in cell biology: How can we understand them better? Molecular Biology of the Cell. 2014;**25**(12):1819-1823. DOI: 10.1091/

[12] Parton DL, Klingelhoefer JW, Sansom MS. Aggregation of model membrane proteins, modulated by hydrophobic mismatch, membrane curvature, and protein class. Biophysical Journal. 2011;**101**(3):691-699. DOI:

[13] Simons K, Toomre D. Lipids rafts and signal transduction. Nature. 2000;**1**(1):31-39. DOI: 10.1038/35036052

[15] Bannenberg GL, Chiang N, Ariel A, Arita M, Tjonahen E, Gotlinger KH, et al. Molecular circuits of resolution: Formation and actions of resolvins and protectins. The Journal of Immunology. 2005;**174**(7):4345-4355. DOI: 10.4049/

[16] Calder PC. Functional roles of fatty acids and their effects on human health. Journal of Parenteral and Enteral Nutrition. 2015;**39**(1 Suppl):18S-32S. DOI: 10.1177/0148607115595980

10.1016/j.bpj.2011.06.048

[14] Waitzberg DL, Torrinhas RS. Fish oil lipid emulsions and immune response? What clinicians need to know. Nutrition in Clinical Practice. 2009;**24**(4):487-499. DOI:

10.1177/0884533609339071

jimmunol.174.7.4345

1999;**2**(2):105-107

mbc.E13-09-0516

*DOI: http://dx.doi.org/10.5772/intechopen.81523*

[Internet]. 2013. Available from: http:// lipidlibrary.aocs.org/Primer/content. cfm?ItemNumber=39371&navItemNum ber=19200 [Accessed: 19 May 2017]

[2] Dewich PM. Medicional Natural Products—A Biosynthetic Approach. 2nd ed. Chichester: John Wiley & Sons;

[3] Fahy E, Subramanian S, Brown HA, Glass CK, Merrill AH, Murphy RC, et al. A comprehensive classification system for lipids. Journal of Lipid Research. 2005;**46**(5):839-862. DOI: 10.1194/jlr.

[4] Quehenberger O, Armando AM, Brown AH, Milne SB, Myers DS, Merril AH, et al. Lipidomics reveals a remarkable diversity of lipids in human plasma. Journal of Lipid Research. 2010;**51**(11):3299-3305. DOI: 10.1194/jlr.

[5] Han X. Lipidomics for studying metabolism. Nature Reviews

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php [Accessed: 19 May 2017]

[7] Brugger B. Analysis of the lipid composition of cells and subcellular organelles by eletrospray ionization mass spectrometry. Annual Review of Biochemistry. 2014;**83**:79-98. DOI: 10.1146/ annurev-biochem-060713-035324

[8] Rolim AEH, Henrique-Araújo R, Ferraz EG, Dultra FKAA, Fernandez LG.

2015;**554**(2):131-139. DOI: 10.1016/j.

Lipidomics in the study of lipid metabolism: Current perspectives in theomic sciences. Gene.

Endocrinology. 2016;**12**(11):668-679.

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*Analytical Tools for Lipid Assessment in Biological Assays DOI: http://dx.doi.org/10.5772/intechopen.81523*

## **References**

*Advances in Lipid Metabolism*

**34**

**Author details**

and Ljubica Tasic3

Sao Paulo, Brazil

Banny Silva Barbosa Correia1

\*

Paulo (USP), School of Medicine, Sao Paulo, Brazil

4 Agilent Technologies Brasil Ltda., Barueri, Sao Paulo, Brazil

\*Address all correspondence to: ljubica@iqm.unicamp.br

provided the original work is properly cited.

© 2018 The Author(s). Licensee IntechOpen. 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,

, Raquel Susana Torrinhas2

1 Chemistry Institute, University of Sao Paulo (USP), Sao Carlos, Sao Paulo, Brazil

2 Department of Gastroenterology, Surgical Division (LIM 35), University of Sao

3 Chemistry Institute, Campinas State University (UNICAMP), Campinas,

, William Yutaka Ohashi3,4

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[50] TIB L, Geloneze B, Pareja JC, Calixto AR, Ferreira MMC, Marsaioli AJ. Omics prospective monitoring of bariatric surgery: Roux-En-Y gastric bypass outcomes using mixed-resolved 1H NMR-based metabolomics. Journal of Integrative Biology. 2016;**20**(7):415- 423. DOI: 10.1089/omi.2016.0061

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[54] Sethi S, Hayashi MAF, Barbosa BS, Pontes JGM, Tasic L, Brietzke E. Lipidomics, biomarkers, and schizophrenia: A current perspective. In: Sussulini A, editor. Metabolomics: From Fundamentals to Clinical Applications, Advances in Experimental Medicine and Biology. Charm: Springer; 2017. pp. 265-290. DOI: 10.1007/978-3-319-47656-8\_11

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[56] Barrilero R, Gil M, Amigo N, Dias CB, Wood LG, Garg ML, et al. LipSpin: A new bioinformatics tool for quantitative 1H NMR lipid profiling. Analytical Chemistry. 2018;**90**(3):2031-2040. DOI: 10.1021/ acs.analchem.7b04148

[57] Fernando H, Bhopale KK, Kondraganti S, Kaphalia BS, Ansari GAS. Lipidomic changes in rat liver after long-term exposure to ethanol. Toxicology and Applied Pharmacology. 2011;**255**(2):127-137. DOI: 10.1016/j. taap.2011.05.022

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[60] Meikle PJ, Wong G, Barlow CK, Weir JM, Greeve MA, MacIntosh GL, et al. Plasma lipid profiling shows similar associations with Prediabetes and type 2 diabetes. PLoS One. 2013;**8**(9):e-74341. DOI: 10.1371/

journal.pone.0074341

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neurobiolaging.2013.08.001

10.1002/mrm.21374

liv.13451

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Boesiger P. SELOVS: Brain MRSI localization based on highly selective T1- and B1-insensitive outer-volume suppression at 3T. Magnetic Resonance in Medicine. 2008;**59**(1):40-51. DOI: 10.1002/mrm.21374

*Advances in Lipid Metabolism*

[46] Zhang Y, Zhao Y, Shen G, Zhong S, Feng J. NMR spectroscopy in

analysis for distinguishing plant origin of edible oils. Journal of Food Composition and Analysis. 2018;**69**:140- 148. DOI: 10.1016/j.jfca.2018.03.006

[47] Vafaeyan H, Ebrahimzadeh SA, Rahimian N, Alavijeh SK, Madadi A, Faeghi F, et al. Quantification of diagnostic biomarkers to detect multiple sclerosis lesions employing 1H-MRSI at 3T. Australasian Physical & Engineering Sciences in Medicine. 2015;**38**(4):611- 618. DOI: 10.1007/s13246-015-0390-1

[48] Dyrby M, Petersen M, Whittaker AK, Lambert L, Nørgaard L, Bro R, et al. Analysis of lipoproteins using 2D diffusion-edited NMR spectroscopy and multi-way chemometrics. Analytica Chimica Acta. 2005;**531**(2):209-216. DOI: 10.1016/j.aca.2004.10.052

[49] Kostara CE, Papathanasiou A, Psychogios N, Cung MT, Elisaf MS, Goudevenos J, et al. NMR-based lipidomic analysis of blood lipoproteins

differentiates the progression of coronary heart disease. Journal of Proteome Research. 2014;**13**(5):2585- 2598. DOI: 10.1021/pr500061n

[50] TIB L, Geloneze B, Pareja JC, Calixto AR, Ferreira MMC, Marsaioli AJ. Omics prospective monitoring of bariatric surgery: Roux-En-Y gastric bypass outcomes using mixed-resolved 1H NMR-based metabolomics. Journal of Integrative Biology. 2016;**20**(7):415- 423. DOI: 10.1089/omi.2016.0061

[51] Barbosa BS, Martins LG, TBBC C, Cruz G, Tasic L. Qualitative and quantitative NMR approaches in blood serum lipidomics. In: Guest P, editor. Investigation of Early Nutrition effects on Long-Term Health—Methods in Molecular Biology. New York: Humana Press; 2018. pp. 365-379. DOI:

10.1007/978-1-4939-7614-0\_25

conjugation with multivariate statistical

[52] Gross RW, Han X. Lipidomics at the interface of structure and function in systems biology. Cell. 2011;**18**(3):284-291. DOI: 10.1016/j.

[53] Sethi S, Hayashi M, Sussulini A, Tasic L, Brietzke E. Analytical approaches for lipidomics and its potential applications in neuropsychiatric disorders. The World Journal of Biological Psychiatry. 2017;**18**(7):506-520. DOI: 10.3109/15622975.2015.1117656

[54] Sethi S, Hayashi MAF, Barbosa BS, Pontes JGM, Tasic L, Brietzke E. Lipidomics, biomarkers, and

schizophrenia: A current perspective. In: Sussulini A, editor. Metabolomics: From Fundamentals to Clinical

Applications, Advances in Experimental

Medicine and Biology. Charm: Springer; 2017. pp. 265-290. DOI: 10.1007/978-3-319-47656-8\_11

[55] Gallo V, Intini N, Mastrorilli P, Latronico M, Scapicchio P, Triggiani M, et al. Performance assessment in fingerprinting and multi component quantitative NMR analyses. Analytical Chemistry. 2015;**87**(13):6709-6717. DOI:

10.1021/acs.analchem.5b00919

acs.analchem.7b04148

taap.2011.05.022

[57] Fernando H, Bhopale KK, Kondraganti S, Kaphalia BS, Ansari GAS. Lipidomic changes in rat liver after long-term exposure to ethanol. Toxicology and Applied Pharmacology. 2011;**255**(2):127-137. DOI: 10.1016/j.

[58] Henning A, Schar M, Schulfe RF, Wilm B, Pruessmann KP,

[56] Barrilero R, Gil M, Amigo N, Dias CB, Wood LG, Garg ML, et al. LipSpin: A new bioinformatics tool for quantitative 1H NMR lipid profiling. Analytical Chemistry. 2018;**90**(3):2031-2040. DOI: 10.1021/

chembiol.2011.01.014

**38**

[59] Traussnigg S, Kienbacher C, Gajdošík M, Valkovič L, Halilbasic E, Stiff J, et al. Ultra-high-field magnetic resonance spectroscopy in non-alcoholic fatty liver disease: Novel mechanistic and diagnostic insights of energy metabolism in nonalcoholic steatohepatitis and advanced fibrosis. Metabolic Liver Disease. 2017;**37**(10):1544-1553. DOI: 10.1111/ liv.13451

[60] Meikle PJ, Wong G, Barlow CK, Weir JM, Greeve MA, MacIntosh GL, et al. Plasma lipid profiling shows similar associations with Prediabetes and type 2 diabetes. PLoS One. 2013;**8**(9):e-74341. DOI: 10.1371/ journal.pone.0074341

[61] Whiley L, Sen A, Heaton J, Proitsi P, García-Gómez D, Leung R, et al. Evidence of altered phosphatidylcholine metabolism in Alzheimer's disease. Neurobiology of Aging. 2013;**35**(2):1-8. DOI: 10.1016/j. neurobiolaging.2013.08.001

**41**

atoms [6].

**Chapter 3**

*Fatiha AID*

**Abstract**

**1. Introduction**

Plant Lipid Metabolism

acids can have industrial and/or pharmaceutical applications.

other organisms and highlighting the specificity of plants.

acid having a trans-type double bond: Δ3 16: 1*t* [5].

**Keywords:** fatty acid, lipids, biosynthesis, plant

In plants, the synthesis of fatty acids takes place in the chloroplast and the fatty acid synthase is prokaryotic type. In plants, the structure of membrane lipids is different from that of eukaryotic cells. The membranes of the chloroplasts are essentially formed of galatolipids. This chapter will also focus on the structure and biosynthesis of fatty acids and membrane lipids in plants. Lipids of seeds are essentially composed of TAG; it would be interesting to describe their synthesis during the maturation of the seeds. Some plants contain in their reserve lipids unconventional fatty acids such as gamma linolenic acid in *Borrago officinalis* L., short-chain fatty acids C: 12 and C: 10, fatty acids with very long chains, and fatty acids that are cyclical. All of these fatty

Plants produce the majority of lipids in the world. These lipids are the main source of calories and essential fatty acids for men and animals. Plants synthesize a huge variety of fatty acids although only a few are major and common constituents [1] like palmitic, oleic, linoleic, and linolenic acids. Like other eukaryotes, lipids are necessary for the biogenesis of cell membranes, as signal molecules and especially as a source of carbon and energy. In plants, carbon, energy, and reducing power are needed for fatty acid biosynthesis derived from photosynthesis in chloroplasts [2]. This chapter will describe lipid biosynthesis in plants by signaling differences with

Fatty acid biosynthesis in plants occurs in the chloroplasts of green tissue and in the plastids of nonphotosynthetic tissues and not in the cytosol as in the animal cell. Although *de novo* synthesis is located in the stroma, plant mitochondria are capable of limited fatty acid synthesis [3]. The plastid membranes are mainly composed of galactolipids, while those of extrachloroplast membranes consist of phospholipids as in the animal cell [4]. Fatty acids in cell membranes consist mainly of palmitic, stearic, oleic, linoleic, and linolenic acids. All double bonds are of cis type. However, in the chloroplast, phosphatidyl glycerol (PG) is acylated with an unusual

Photosynthetic tissues of higher plants contain 60–70% trienoic fatty acids. The so-called "C18: 3" plants are generally the most advanced families of angiosperms (pea, spinach, etc.) whose position *sn-*2 of the galactolipids is esterified exclusively by polyunsaturated fatty acids with 18 carbon atoms. The "C16: 3" plants are generally the less evolved families of angiosperms (Brassicaceae) whose position *sn-*2 of the galactolipids is esterified by polyunsaturated fatty acids with 16 or 18 carbon

## **Chapter 3** Plant Lipid Metabolism

*Fatiha AID*

## **Abstract**

In plants, the synthesis of fatty acids takes place in the chloroplast and the fatty acid synthase is prokaryotic type. In plants, the structure of membrane lipids is different from that of eukaryotic cells. The membranes of the chloroplasts are essentially formed of galatolipids. This chapter will also focus on the structure and biosynthesis of fatty acids and membrane lipids in plants. Lipids of seeds are essentially composed of TAG; it would be interesting to describe their synthesis during the maturation of the seeds. Some plants contain in their reserve lipids unconventional fatty acids such as gamma linolenic acid in *Borrago officinalis* L., short-chain fatty acids C: 12 and C: 10, fatty acids with very long chains, and fatty acids that are cyclical. All of these fatty acids can have industrial and/or pharmaceutical applications.

**Keywords:** fatty acid, lipids, biosynthesis, plant

## **1. Introduction**

Plants produce the majority of lipids in the world. These lipids are the main source of calories and essential fatty acids for men and animals. Plants synthesize a huge variety of fatty acids although only a few are major and common constituents [1] like palmitic, oleic, linoleic, and linolenic acids. Like other eukaryotes, lipids are necessary for the biogenesis of cell membranes, as signal molecules and especially as a source of carbon and energy. In plants, carbon, energy, and reducing power are needed for fatty acid biosynthesis derived from photosynthesis in chloroplasts [2]. This chapter will describe lipid biosynthesis in plants by signaling differences with other organisms and highlighting the specificity of plants.

Fatty acid biosynthesis in plants occurs in the chloroplasts of green tissue and in the plastids of nonphotosynthetic tissues and not in the cytosol as in the animal cell. Although *de novo* synthesis is located in the stroma, plant mitochondria are capable of limited fatty acid synthesis [3]. The plastid membranes are mainly composed of galactolipids, while those of extrachloroplast membranes consist of phospholipids as in the animal cell [4]. Fatty acids in cell membranes consist mainly of palmitic, stearic, oleic, linoleic, and linolenic acids. All double bonds are of cis type. However, in the chloroplast, phosphatidyl glycerol (PG) is acylated with an unusual acid having a trans-type double bond: Δ3 16: 1*t* [5].

Photosynthetic tissues of higher plants contain 60–70% trienoic fatty acids. The so-called "C18: 3" plants are generally the most advanced families of angiosperms (pea, spinach, etc.) whose position *sn-*2 of the galactolipids is esterified exclusively by polyunsaturated fatty acids with 18 carbon atoms. The "C16: 3" plants are generally the less evolved families of angiosperms (Brassicaceae) whose position *sn-*2 of the galactolipids is esterified by polyunsaturated fatty acids with 16 or 18 carbon atoms [6].

Plant lipids have a substantial impact on the world economy and human nutrition. The majority of oils used by humans are triacylglycerols derived from seeds or fruits. Indeed, the seeds are subdivided into three categories according to their reserve. Seeds that contain more than 45% protein are called protein seeds. Starchy seeds contain more than 70% of carbohydrates like cereals. Oleaginous seeds contain more than 50% of lipids in the form of triacylglycerol esterified generally by palmitic, oleic, linoleic, and linolenic acids in majority of seeds. Some plants can produce unusual fatty acids like hydroxyl fatty acids, cyclopropane fatty acids, epoxy fatty acids, and conjugated unsaturated fatty acids in their seed oils, many of which have useful industrial applications [7]. These unusual fatty acids accumulate preferentially in triacylglycerols.

## **2. Fatty acid synthesis**

In plants, *de novo* fatty acid biosynthesis mainly takes place in the plastidial compartment [8] from acetyl CoA, which is a direct product of photosynthesis. Plastid pyruvate dehydrogenase (EC 1.2.4.1) is the main route for a rapid and stable supply of acetyl CoA through its action on pyruvate (resulting from glycolysis or the pentose phosphate pathway). Another possible source is the import and activation of free acetate by acetyl CoA synthase (ACS, EC 6.2.1.1) [9]. The major product of FAS is palmitic acid, except the elongation of palmitic acid and the desaturation of stearic acid which take place in the chloroplast. Other changes (elongation, desaturation, hydroxylation, and epoxidation) occur mainly in the endoplasmic reticulum.

Two enzyme systems are required for fatty acid formation: acetyl CoA carboxylase (ACCase, EC 6.4.1.2) of which two forms have been identified in plants [10] and fatty acid synthase which is a multienzyme complex present in the stroma of chloroplasts [11].

## **2.1 Acetyl CoA carboxylase (ACCase)**

The first enzyme complex is the ACCase that catalyzes an ATP-dependent carboxylation of acetyl CoA to malonyl CoA. For plants, acetyl CoA carboxylase (ACCase) directs the flow of carbon from photosynthesis to primary and secondary metabolites. Two distinct molecular forms of ACCase have been identified, a multiprotein complex and a multifunctional protein [12] (**Figure 1**).

#### **Figure 1.**

*Structure of the two types of ACCase. (A) The multisubunit (MS complex) ACCase and (B) the multifunctional (MF) ACCase. BCCP, biotin carboxyl carrier protein; BC, biotin carboxylase; α and β CT, α and β carboxy transferase; VLCFA, very long-chain fatty acids.*

**43**

*Plant Lipid Metabolism*

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

acetyl CoA to form malonyl CoA.

**2.2 Fatty acid synthase (FAS)**

fatty acids are in acyl CoA form [15].

pathway are acyl-ACPs.

two-carbon addition (**Figure 2**).

(16:0-ACP) to stearoyl-ACP (18:0-ACP).

sensitive (MF), in soybean leaf chloroplasts [14].

The multisubunit (MS complex) ACCase, present in plastids of all plants, except *Poaceae* and *Geraniaceae*, is involved in *de novo* fatty acid synthesis [13]. It is composed of four independent polypeptides: biotin carboxyl carrier protein (BCCP), biotin carboxylase (BC) and α and β carboxy transferase (α and β CT). The biotin carboxylase (BC) subunit catalyzes the ATP-dependent carboxylation of the biotinyl moeity on biotin carboxyl carrier protein (BCCP), and the carboxytransferase (CT) subunits catalyze the transfer of activated carboxyl groups from BCCP to

The multifunctional (MF) ACCase, consisting of a single 220–240 kDa polypeptide with BCCP, BC, and CT domains, is nuclear encoded except the αCT subunit which is encoded by the plastidial genome [13]. In all plants, MF ACCase is involved

The sensitivity of plastidial ACCase to sethoxydim and the presence of a 220 kDa biotinylated polypeptide in soybean plastids provide a biochemical indication for the possible presence of two ACCase isoforms, one resistant (MS) and one

The second enzyme complex involved in *de novo* synthesis is the fatty acid synthase (FAS). In nature, fatty acid synthases are subdivided into two groups. The fatty acid synthase type I which is characterized by a large, multifunctional proteins typical of yeast and mammals and the fatty acid synthase type II, found in prokaryotes which is composed of four dissociable proteins that catalyze individual reactions. Although plant cells are eukaryotic, the fatty acid synthase found in plastids is of type II [15]. Plant fatty acid synthase has inherited from photosynthetic prokaryotes; plastids being considered by the endosymbiotic theory as an old cyanobacteria. In plants, acyl carrier protein (ACP) is used as the acyl carrier for the various intermediate for fatty acid synthase unlike other eukaryotic cells where

The initial substrates for fatty acid biosynthesis are acetyl CoA and malonyl-ACPs. The transfer of malonyl moiety from CoA to ACP is catalyzed by malonyl CoA:ACP transacylase (MAT). After the initial condensation of acetyl CoA and malonyl-ACP, all the intermediates for each step of the fatty acid biosynthetic

AGS is composed of four enzymes: ketoacyl-ACP synthase (KAS, EC 2.3.1.41),

The first condensation takes place between acetyl CoA and malonyl-ACP This reaction is catalyzed by 1,3-ketoacyl-ACP synthase III (KAS, EC 2.3.1.41), one of three ketoacyl synthases in plant systems [15]. KAS I is responsible for the condensations in each elongation cycle up through that producing palmitoyl-ACP (16:0- ACP). KAS II is dedicated to the final plastidial elongation, that of palmitoyl-ACP

The β-ketoacyl-ACP formed during the condensation reaction successively undergoes a reduction reaction by β-ketoacyl-ACP reductase (EC 1.1.1.100), dehydration by the β hydroxyacyl-ACP dehydratase (EC 4.2.1.59) and a further reduction by enoylacyl-ACP reductase (EC 1.3.1.9) to give butyryl-ACP. The coenzyme of the two oxidation-reduction reactions is NADPH (**Figure 3**). The butyryl-ACP formed

β-ketoacyl-ACP reductase (EC 1.1.1.100), hydroxy acyl-ACP dehydrase (EC 4.2.1.59), and enoylacyl-ACP reductase (EC 1.3.1.9). All components of fatty acid synthase occur in plastids, although they are encoded in the nuclear genome and synthesized on cytosolic ribosomes. There are four sequential reactions involved in

in very long-chain fatty acid and flavonoid biosynthesis in the cytosol [13].

#### *Plant Lipid Metabolism DOI: http://dx.doi.org/10.5772/intechopen.81355*

*Advances in Lipid Metabolism*

**2. Fatty acid synthesis**

chloroplasts [11].

**2.1 Acetyl CoA carboxylase (ACCase)**

**42**

**Figure 1.**

*Structure of the two types of ACCase. (A) The multisubunit (MS complex) ACCase and (B) the* 

*and β carboxy transferase; VLCFA, very long-chain fatty acids.*

*multifunctional (MF) ACCase. BCCP, biotin carboxyl carrier protein; BC, biotin carboxylase; α and β CT, α*

Plant lipids have a substantial impact on the world economy and human nutrition. The majority of oils used by humans are triacylglycerols derived from seeds or fruits. Indeed, the seeds are subdivided into three categories according to their reserve. Seeds that contain more than 45% protein are called protein seeds. Starchy seeds contain more than 70% of carbohydrates like cereals. Oleaginous seeds contain more than 50% of lipids in the form of triacylglycerol esterified generally by palmitic, oleic, linoleic, and linolenic acids in majority of seeds. Some plants can produce unusual fatty acids like hydroxyl fatty acids, cyclopropane fatty acids, epoxy fatty acids, and conjugated unsaturated fatty acids in their seed oils, many of which have useful industrial applications [7]. These unusual fatty acids accumulate preferentially in triacylglycerols.

In plants, *de novo* fatty acid biosynthesis mainly takes place in the plastidial compartment [8] from acetyl CoA, which is a direct product of photosynthesis. Plastid pyruvate dehydrogenase (EC 1.2.4.1) is the main route for a rapid and stable supply of acetyl CoA through its action on pyruvate (resulting from glycolysis or the pentose phosphate pathway). Another possible source is the import and activation of free acetate by acetyl CoA synthase (ACS, EC 6.2.1.1) [9]. The major product of FAS is palmitic acid, except the elongation of palmitic acid and the desaturation of stearic acid which take place in the chloroplast. Other changes (elongation, desaturation, hydroxylation, and epoxidation) occur mainly in the endoplasmic reticulum. Two enzyme systems are required for fatty acid formation: acetyl CoA carboxylase (ACCase, EC 6.4.1.2) of which two forms have been identified in plants [10] and fatty acid synthase which is a multienzyme complex present in the stroma of

The first enzyme complex is the ACCase that catalyzes an ATP-dependent carboxylation of acetyl CoA to malonyl CoA. For plants, acetyl CoA carboxylase (ACCase) directs the flow of carbon from photosynthesis to primary and secondary metabolites. Two distinct molecular forms of ACCase have been identified, a

multiprotein complex and a multifunctional protein [12] (**Figure 1**).

The multisubunit (MS complex) ACCase, present in plastids of all plants, except *Poaceae* and *Geraniaceae*, is involved in *de novo* fatty acid synthesis [13]. It is composed of four independent polypeptides: biotin carboxyl carrier protein (BCCP), biotin carboxylase (BC) and α and β carboxy transferase (α and β CT). The biotin carboxylase (BC) subunit catalyzes the ATP-dependent carboxylation of the biotinyl moeity on biotin carboxyl carrier protein (BCCP), and the carboxytransferase (CT) subunits catalyze the transfer of activated carboxyl groups from BCCP to acetyl CoA to form malonyl CoA.

The multifunctional (MF) ACCase, consisting of a single 220–240 kDa polypeptide with BCCP, BC, and CT domains, is nuclear encoded except the αCT subunit which is encoded by the plastidial genome [13]. In all plants, MF ACCase is involved in very long-chain fatty acid and flavonoid biosynthesis in the cytosol [13].

The sensitivity of plastidial ACCase to sethoxydim and the presence of a 220 kDa biotinylated polypeptide in soybean plastids provide a biochemical indication for the possible presence of two ACCase isoforms, one resistant (MS) and one sensitive (MF), in soybean leaf chloroplasts [14].

## **2.2 Fatty acid synthase (FAS)**

The second enzyme complex involved in *de novo* synthesis is the fatty acid synthase (FAS). In nature, fatty acid synthases are subdivided into two groups. The fatty acid synthase type I which is characterized by a large, multifunctional proteins typical of yeast and mammals and the fatty acid synthase type II, found in prokaryotes which is composed of four dissociable proteins that catalyze individual reactions. Although plant cells are eukaryotic, the fatty acid synthase found in plastids is of type II [15]. Plant fatty acid synthase has inherited from photosynthetic prokaryotes; plastids being considered by the endosymbiotic theory as an old cyanobacteria. In plants, acyl carrier protein (ACP) is used as the acyl carrier for the various intermediate for fatty acid synthase unlike other eukaryotic cells where fatty acids are in acyl CoA form [15].

The initial substrates for fatty acid biosynthesis are acetyl CoA and malonyl-ACPs. The transfer of malonyl moiety from CoA to ACP is catalyzed by malonyl CoA:ACP transacylase (MAT). After the initial condensation of acetyl CoA and malonyl-ACP, all the intermediates for each step of the fatty acid biosynthetic pathway are acyl-ACPs.

AGS is composed of four enzymes: ketoacyl-ACP synthase (KAS, EC 2.3.1.41), β-ketoacyl-ACP reductase (EC 1.1.1.100), hydroxy acyl-ACP dehydrase (EC 4.2.1.59), and enoylacyl-ACP reductase (EC 1.3.1.9). All components of fatty acid synthase occur in plastids, although they are encoded in the nuclear genome and synthesized on cytosolic ribosomes. There are four sequential reactions involved in two-carbon addition (**Figure 2**).

The first condensation takes place between acetyl CoA and malonyl-ACP This reaction is catalyzed by 1,3-ketoacyl-ACP synthase III (KAS, EC 2.3.1.41), one of three ketoacyl synthases in plant systems [15]. KAS I is responsible for the condensations in each elongation cycle up through that producing palmitoyl-ACP (16:0- ACP). KAS II is dedicated to the final plastidial elongation, that of palmitoyl-ACP (16:0-ACP) to stearoyl-ACP (18:0-ACP).

The β-ketoacyl-ACP formed during the condensation reaction successively undergoes a reduction reaction by β-ketoacyl-ACP reductase (EC 1.1.1.100), dehydration by the β hydroxyacyl-ACP dehydratase (EC 4.2.1.59) and a further reduction by enoylacyl-ACP reductase (EC 1.3.1.9) to give butyryl-ACP. The coenzyme of the two oxidation-reduction reactions is NADPH (**Figure 3**). The butyryl-ACP formed

#### **Figure 2.**

*Plant fatty acid biosynthesis. This chain requires a carboxylation reaction of acetyl CoA to malonyl CoA, an activation reaction of malonyl CoA to malonyl-ACP, a condensation reaction between acetyl CoA and malonyl-ACP to form β-ketoacyl-ACP, which undergoes in turn a reduction reaction, dehydration, and a second reduction extending the fatty acid of two carbon atoms.*

will be extended by two further C2 units after further condensation with malonyl-ACP. The β-ketoacyl-ACP synthase I (KASI) catalyzes this reaction. After seven rounds of cycle, palmitoyl-ACP is formed.

Although the final product of fatty acid synthase is palmitic acid, two other common fatty acids are synthesized in the chloroplast stroma. These are stearic and oleic acids. The palmitoyl-ACP (C16:0-ACP) will be extended by two new units to form a stearoyl-ACP (C18:0-ACP) chain by a plastid soluble stearoyl-ACP synthase which is a multienzymatic complex composed of four enzymes (KASII, enoyl-ACP reductase, hydroxyacyl-ACP dehydrase, and enoylacyl-ACP reductase) [16].

The formed stearoyl-ACP is then desaturated with a plastidial soluble stearoyl-ACP desaturase (SAD, EC 1.14. 19.2) in oleoyl-ACP (C18:1Δ9-ACP) [17]. This enzyme is a nuclear-encoded, plastid-localized soluble desaturase that introduces the first Δ9 double bond into the saturated fatty acid resulting in the conversion of 18:0-ACP into 18:1Δ9-ACP [18].

The lack of structural similarity between plant and mammalian desaturase reflects the facts that the fatty acid substrates are on different carriers (ACP and

**45**

the environment.

**3.1 Importance of thioesterases**

*Plant Lipid Metabolism*

**Figure 3.**

*stearoyl-ACP desaturase.*

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

CoA, respectively),that the enzymes utilize different electron donors (ferredoxin vs. cytochrome b5), and that the plant enzyme is soluble, whereas the animal and fungal enzymes are integral membrane protein [19]. This enzyme plays a key role in

*Schematic representation of the export of fatty acids from the plastid to the cytosol. ACC, acetyl CoA carboxylase; ACP, acyl carrier protein; FA, fatty acid; CoA, coenzyme A; FAS, fatty acid synthase; FAT A/B, fatty acyl-ACP thioesterase A/B; LACS, long-chain acyl CoA synthase; PAE, palmitoyl-ACP elongase; SAD,* 

In addition to the soluble acyl-ACP desaturases, the fatty acids synthesized in the chloroplast (palmitate, stearate, and oleate) are desaturated by membrane-bound desaturases that utilize complex lipid substrates such as phosphatidylcholine (PC) in the endoplasmic reticulum (ER) or monogalactosyl-diacylglycerol (MGDG) in the plastid [16]. These fatty acids were used for the synthesis of glycerolipids by two distinct metabolic pathways (prokaryotic and eukaryotic pathway) and in different

The importance of both biosynthetic pathway depends on the plant species. In the "C18:3" plants photosynthetically active, only the extrachloroplast galactolipid pathway is functional; in the case of "C16:3" plants, the two pathways coexist and their importance differs according to the species and the conditions of

The flow of fatty acids (palmitoyl-ACP, oleoyl-ACP, and to a lesser extent stearoyl-ACP) through the two pathways would be subject to severe control. Two acyl-ACPs thioesterase enzymes and a chloroplast glycerol 3-phosphate acyl transferase play a very important role. Indeed, the acyl residues enter directly into the extrachloroplast pathway after having been hydrolyzed by fatty acyl-ACPs thioesterases FAT (A EC 3.1.2.14 and B EC 3.1.2.22) [22, 23] or in the chloroplast pathway

The released palmitic, oleic, and stearic acids are then activated into coenzyme A ester by the action of long-chain acyl-CoA synthetase (LACS, EC 6.2.1.3) [24] and

determining the ratio of saturated to unsaturated fatty acids [17].

**3. Glycerolipids as substrates for desaturation**

cellular compartments (plastids and ER) [20].

after being acylated by acyl transferases [21].

are exported to the cytosol (**Figure 3**).

#### **Figure 3.**

*Advances in Lipid Metabolism*

**44**

**Figure 2.**

will be extended by two further C2 units after further condensation with malonyl-ACP. The β-ketoacyl-ACP synthase I (KASI) catalyzes this reaction. After seven

*Plant fatty acid biosynthesis. This chain requires a carboxylation reaction of acetyl CoA to malonyl CoA, an activation reaction of malonyl CoA to malonyl-ACP, a condensation reaction between acetyl CoA and malonyl-ACP to form β-ketoacyl-ACP, which undergoes in turn a reduction reaction, dehydration, and a* 

Although the final product of fatty acid synthase is palmitic acid, two other common fatty acids are synthesized in the chloroplast stroma. These are stearic and oleic acids. The palmitoyl-ACP (C16:0-ACP) will be extended by two new units to form a stearoyl-ACP (C18:0-ACP) chain by a plastid soluble stearoyl-ACP synthase which is a multienzymatic complex composed of four enzymes (KASII, enoyl-ACP reductase, hydroxyacyl-ACP dehydrase, and enoylacyl-ACP reductase) [16].

The formed stearoyl-ACP is then desaturated with a plastidial soluble stearoyl-

ACP desaturase (SAD, EC 1.14. 19.2) in oleoyl-ACP (C18:1Δ9-ACP) [17]. This enzyme is a nuclear-encoded, plastid-localized soluble desaturase that introduces the first Δ9 double bond into the saturated fatty acid resulting in the conversion of

The lack of structural similarity between plant and mammalian desaturase reflects the facts that the fatty acid substrates are on different carriers (ACP and

rounds of cycle, palmitoyl-ACP is formed.

*second reduction extending the fatty acid of two carbon atoms.*

18:0-ACP into 18:1Δ9-ACP [18].

*Schematic representation of the export of fatty acids from the plastid to the cytosol. ACC, acetyl CoA carboxylase; ACP, acyl carrier protein; FA, fatty acid; CoA, coenzyme A; FAS, fatty acid synthase; FAT A/B, fatty acyl-ACP thioesterase A/B; LACS, long-chain acyl CoA synthase; PAE, palmitoyl-ACP elongase; SAD, stearoyl-ACP desaturase.*

CoA, respectively),that the enzymes utilize different electron donors (ferredoxin vs. cytochrome b5), and that the plant enzyme is soluble, whereas the animal and fungal enzymes are integral membrane protein [19]. This enzyme plays a key role in determining the ratio of saturated to unsaturated fatty acids [17].

### **3. Glycerolipids as substrates for desaturation**

In addition to the soluble acyl-ACP desaturases, the fatty acids synthesized in the chloroplast (palmitate, stearate, and oleate) are desaturated by membrane-bound desaturases that utilize complex lipid substrates such as phosphatidylcholine (PC) in the endoplasmic reticulum (ER) or monogalactosyl-diacylglycerol (MGDG) in the plastid [16]. These fatty acids were used for the synthesis of glycerolipids by two distinct metabolic pathways (prokaryotic and eukaryotic pathway) and in different cellular compartments (plastids and ER) [20].

The importance of both biosynthetic pathway depends on the plant species. In the "C18:3" plants photosynthetically active, only the extrachloroplast galactolipid pathway is functional; in the case of "C16:3" plants, the two pathways coexist and their importance differs according to the species and the conditions of the environment.

#### **3.1 Importance of thioesterases**

The flow of fatty acids (palmitoyl-ACP, oleoyl-ACP, and to a lesser extent stearoyl-ACP) through the two pathways would be subject to severe control. Two acyl-ACPs thioesterase enzymes and a chloroplast glycerol 3-phosphate acyl transferase play a very important role. Indeed, the acyl residues enter directly into the extrachloroplast pathway after having been hydrolyzed by fatty acyl-ACPs thioesterases FAT (A EC 3.1.2.14 and B EC 3.1.2.22) [22, 23] or in the chloroplast pathway after being acylated by acyl transferases [21].

The released palmitic, oleic, and stearic acids are then activated into coenzyme A ester by the action of long-chain acyl-CoA synthetase (LACS, EC 6.2.1.3) [24] and are exported to the cytosol (**Figure 3**).

Plants export sufficient fatty acid (16: 0-CoA, 18: 0-CoA, 18: 1-CoA) for lipid synthesis of extraplastid membranes and TAG of seed lipids of all plants.

#### **3.2 Prokaryotic pathway**

The prokaryotic pathway uses acyl-ACPs for PA and PG synthesis in all plants, and galactolipids (MGDG, DGDG, and SQDG) of so-called "C16:3" plants. This pathway is similar to the pathway demonstrated in photosynthetic prokaryotes [22].

The prokaryotic pathway is distinguished from the eukaryotic one by the presence of C16 fatty acids at the *sn*-2 position of the glycerol backbone. This pathway is characterized by the presence of molecular species 18: 3/16: 3 MGDG [23].

The major molecular species of MGDG synthesized by the prokaryotic pathway generally contain α-linolenic acid (C18:3), exclusively on the *sn*-1 position of glycerol backbone, while the *sn*-2 position is esterified by hexadecatrienoic acid (C16:3), resulting in desaturation of palmitic acid. The prokaryotic pathway, exclusively localized in plastids, therefore requires desaturation steps.

The DAG, precursor of prokaryotic MGDG, is an 18:1/16:0 DAG (**Figure 5**). The first molecular species synthesized by MGDG synthase is 18:1/16 0 MGDG. The palmitoyl residue is desaturated to a *cis*-hexadecenoyl residue by an ω9 desaturase, which is specific for both, the *sn*2 position of the fatty acid on glycerol and lipids (MGDG) [24, 25]. The ω9 desaturase is much more active on the palmitoyl residue in the *sn*2 position of glycerol of the MGDG than on the one located in the position *sn*2 of the DGDG.

The ω6 and ω3 desaturases, respectively, catalyze the desaturation of the monounsaturated acyls (hexadecenoyl and oleoyl) and diunsaturated (hexadecadienoyl and linoleoyl) residues. These desaturases have no specificity with respect to the length of the fatty acid chain or its position on glycerol. The ω6 desaturase acts equally well on the hexadecenoyl and oleoyl residues located at the *sn*2 and *sn*1 positions of the MGDGs and DGDGs. The desaturation of palmitic acid to hexadecenoyl acid is a prerequisite for other desaturations [26] (**Figure 4**).

This desaturation scheme is similar to that proposed for the desaturation of lipid acyls in the blue seaweed *Anabaena variabilis* [22].

Phosphatidyl glycerol synthesis occurs in both "C16:3" and "C18: 3" plants in the chloroplast. It involves in a first step the phosphatidic acid (PA) and CDP DAG which is of prokaryotic type. About 30–40% of the palmitoyl residue at position *sn*-2 of PG is desaturated at carbon 3 to form 3-*trans*-hexadecenoic acid [27]. The structure of this fatty acid is unusual; in plants, all the double bonds of the fatty acids of the membrane lipids are of cis type with the exception of this fatty acid.

#### **3.3 Eukaryotic pathway**

The second pathway, called the "eukaryotic" pathway, leads to the formation of two types of MGDG molecular species; one of them contains α-linoleate in both

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*Plant Lipid Metabolism*

chloroplasts [23, 28].

**Figure 5.**

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

positions of glycerol and the other molecular species contains α-linoleate in position *sn*-2 and palmitate in position *sn*-1. This pathway requires cooperation between plastids and the endoplasmic reticulum for the formation of glyceroglycolipids in

*Scheme of possible ways for the synthesis of eukaryotic-type MGDG in spinach, C16: 3 plants according to [25].*

The oleate integrated into PC molecules at the position *sn*-2 of glycerol backbone undergoes a succession of desaturations catalyzed by the (ω-6, Δ12) oleate desaturase, still identified by the fad2 mutation of *Arabidopsis* and allowing the synthesis of linoleic acid and the (ω-3, Δ15) linoleate desaturase, identified by the fad3 mutation of *Arabidopsis*, which allows the synthesis of α-linoleic acid. Mutants deficient for the lysophosphatidylcholine acyltransferase (LPCAT1 and LPCAT2 genes) have

After desaturation as acyl-PC, a part of them, probably in the form of DAG, returns to the chloroplast and contributes to the formation of chloroplast galactolipids (**Figure 5**). These DAGs can be desaturated by chloroplast desaturases [30, 31]. It is therefore possible to judge the relative contributions of the prokaryotic and eukaryotic pathways by comparing the proportions of eukaryotic 18/18 and 16/18

Membranes of eukaryotic cells have multiple functions in ensuring physical compartmentalization at the cellular and subcellular levels, the regulation of exchanges by the transport of metabolites and macromolecules, cellular communication (hormone receptors, surface antigens, signal transduction, etc.), and in some specific metabolic reactions. In the same cell, it is therefore not surprising to encounter different types of membranes with a specific lipid and protein composition that will determine their respective functions [32]. This synthesis is mainly carried out by two metabolic pathways described as prokaryotic and eukaryotic [33].

Eukaryotic DAGs and prokaryotic DAG structures are the precursors of glycolipid synthesis (SQDG, MGDG, and DGDG). There are therefore two types of glycolipids: prokaryotic glycolipids whose DAG backbone is of the C18/C16 type and which are desaturated exclusively in the plastid and eukaryotic glycolipids including DAGs of the type (C18:1/C18:1 and C16:0/C18:1) are derived from phosphatidylcholine and are desaturated in RE and plastid [34]. The synthesis of glycolipids, being localized in the membranes of the plastid envelope, thus requires a mechanism for importing DAGs of eukaryotic structure. These differences in DAG structure are due to different specificities of the chloroplast and ER acyl transferases. The first step of the prokaryotic pathway is the transfer of the oleate to

reduced levels of polyunsaturated FA (PUFA) in TAGs [29].

glycerolipids with prokaryotic 18/16 glycerolipids.

**4. Synthesis of membrane glycerolipids**

**4.1 Synthesis of plastid lipids**

**Figure 4.** *Possible desaturation scheme of prokaryotic MGDG in plastids.*

#### **Figure 5.**

*Advances in Lipid Metabolism*

**3.2 Prokaryotic pathway**

*sn*2 of the DGDG.

**3.3 Eukaryotic pathway**

Plants export sufficient fatty acid (16: 0-CoA, 18: 0-CoA, 18: 1-CoA) for lipid

The prokaryotic pathway uses acyl-ACPs for PA and PG synthesis in all plants, and galactolipids (MGDG, DGDG, and SQDG) of so-called "C16:3" plants. This pathway is similar to the pathway demonstrated in photosynthetic prokaryotes [22]. The prokaryotic pathway is distinguished from the eukaryotic one by the presence of C16 fatty acids at the *sn*-2 position of the glycerol backbone. This pathway is

The major molecular species of MGDG synthesized by the prokaryotic pathway generally contain α-linolenic acid (C18:3), exclusively on the *sn*-1 position of glycerol backbone, while the *sn*-2 position is esterified by hexadecatrienoic acid (C16:3), resulting in desaturation of palmitic acid. The prokaryotic pathway, exclusively

The DAG, precursor of prokaryotic MGDG, is an 18:1/16:0 DAG (**Figure 5**). The first molecular species synthesized by MGDG synthase is 18:1/16 0 MGDG. The palmitoyl residue is desaturated to a *cis*-hexadecenoyl residue by an ω9 desaturase, which is specific for both, the *sn*2 position of the fatty acid on glycerol and lipids (MGDG) [24, 25]. The ω9 desaturase is much more active on the palmitoyl residue in the *sn*2 position of glycerol of the MGDG than on the one located in the position

The ω6 and ω3 desaturases, respectively, catalyze the desaturation of the monounsaturated acyls (hexadecenoyl and oleoyl) and diunsaturated (hexadecadienoyl and linoleoyl) residues. These desaturases have no specificity with respect to the length of the fatty acid chain or its position on glycerol. The ω6 desaturase acts equally well on the hexadecenoyl and oleoyl residues located at the *sn*2 and *sn*1 positions of the MGDGs and DGDGs. The desaturation of palmitic acid to hexadecenoyl

This desaturation scheme is similar to that proposed for the desaturation of lipid

Phosphatidyl glycerol synthesis occurs in both "C16:3" and "C18: 3" plants in the chloroplast. It involves in a first step the phosphatidic acid (PA) and CDP DAG which is of prokaryotic type. About 30–40% of the palmitoyl residue at position *sn*-2 of PG is desaturated at carbon 3 to form 3-*trans*-hexadecenoic acid [27]. The structure of this fatty acid is unusual; in plants, all the double bonds of the fatty acids of the membrane lipids are of cis type with the exception of this fatty acid.

The second pathway, called the "eukaryotic" pathway, leads to the formation of two types of MGDG molecular species; one of them contains α-linoleate in both

synthesis of extraplastid membranes and TAG of seed lipids of all plants.

characterized by the presence of molecular species 18: 3/16: 3 MGDG [23].

localized in plastids, therefore requires desaturation steps.

acid is a prerequisite for other desaturations [26] (**Figure 4**).

acyls in the blue seaweed *Anabaena variabilis* [22].

**46**

**Figure 4.**

*Possible desaturation scheme of prokaryotic MGDG in plastids.*

*Scheme of possible ways for the synthesis of eukaryotic-type MGDG in spinach, C16: 3 plants according to [25].*

positions of glycerol and the other molecular species contains α-linoleate in position *sn*-2 and palmitate in position *sn*-1. This pathway requires cooperation between plastids and the endoplasmic reticulum for the formation of glyceroglycolipids in chloroplasts [23, 28].

The oleate integrated into PC molecules at the position *sn*-2 of glycerol backbone undergoes a succession of desaturations catalyzed by the (ω-6, Δ12) oleate desaturase, still identified by the fad2 mutation of *Arabidopsis* and allowing the synthesis of linoleic acid and the (ω-3, Δ15) linoleate desaturase, identified by the fad3 mutation of *Arabidopsis*, which allows the synthesis of α-linoleic acid. Mutants deficient for the lysophosphatidylcholine acyltransferase (LPCAT1 and LPCAT2 genes) have reduced levels of polyunsaturated FA (PUFA) in TAGs [29].

After desaturation as acyl-PC, a part of them, probably in the form of DAG, returns to the chloroplast and contributes to the formation of chloroplast galactolipids (**Figure 5**). These DAGs can be desaturated by chloroplast desaturases [30, 31].

It is therefore possible to judge the relative contributions of the prokaryotic and eukaryotic pathways by comparing the proportions of eukaryotic 18/18 and 16/18 glycerolipids with prokaryotic 18/16 glycerolipids.

## **4. Synthesis of membrane glycerolipids**

Membranes of eukaryotic cells have multiple functions in ensuring physical compartmentalization at the cellular and subcellular levels, the regulation of exchanges by the transport of metabolites and macromolecules, cellular communication (hormone receptors, surface antigens, signal transduction, etc.), and in some specific metabolic reactions. In the same cell, it is therefore not surprising to encounter different types of membranes with a specific lipid and protein composition that will determine their respective functions [32]. This synthesis is mainly carried out by two metabolic pathways described as prokaryotic and eukaryotic [33].

#### **4.1 Synthesis of plastid lipids**

Eukaryotic DAGs and prokaryotic DAG structures are the precursors of glycolipid synthesis (SQDG, MGDG, and DGDG). There are therefore two types of glycolipids: prokaryotic glycolipids whose DAG backbone is of the C18/C16 type and which are desaturated exclusively in the plastid and eukaryotic glycolipids including DAGs of the type (C18:1/C18:1 and C16:0/C18:1) are derived from phosphatidylcholine and are desaturated in RE and plastid [34]. The synthesis of glycolipids, being localized in the membranes of the plastid envelope, thus requires a mechanism for importing DAGs of eukaryotic structure. These differences in DAG structure are due to different specificities of the chloroplast and ER acyl transferases. The first step of the prokaryotic pathway is the transfer of the oleate to

#### *Advances in Lipid Metabolism*

a glycerol-3-phosphate at position *sn*-1 by an acyl ACP-glycerol 3 phosphate acyltransferase (EC 2.3.1.1), soluble in the stroma of the plastid [35]. Lysophosphatidic acid (LPA) is thus formed. A second plastid-related plastid acyltransferase, the LPA-ACP acyltransferase, catalyzes the esterification of palmitoyl-ACP at the *sn*-2 position (LPAAT1; EC 2.3.1.51) [36]. This results in the synthesis of 18:1/16:0-PA.

Phosphatidic acid (PA) can either be converted to CDP-DAG by the action of a CTP-phosphatidate cytidylyltransferase (EC 2.7.7.41) which catalyzes the reaction between PA and CTP to form CDP-DAG and pyrophosphate or dephosphorylated to diacylglycerol (DAG) by phosphatidate phosphatase (PAP; EC 3.1.3.4). CDP-DAG will be used for the synthesis of phosphatidyl glycerol (PG) of the plastid [35] and DAG can be used for the synthesis of galactolipids (MGDG, DGDG) or a sulfolipid (sulfoquinovosyldiacylglycerol) (**Figure 6**).

From the phyllogenetic point of view, the difference between so-called "C16:3" and "C18:3" plants is related to the presence of plastid phosphatidate phosphatase in "C16: 3" plants, lost during evolution in "C18:3" plants. The chloroplast enzyme is clearly different from other phosphatidate phosphatases in the cell because it is membrane-bound, strongly associated with the inner membrane of the envelope, has an optimum alkaline pH, and is inhibited by cations such as Mg2+ [37]. The DAG thus produced (18/16 DAG) is at the origin of the glycolipids of prokaryotic structure, SQDG, MGDG, and DGDG (**Figure 5**).

### *4.1.1 Synthesis of monogalactosyl diacylglycerol (MGDG)*

MGDG is synthesized in a single step by a 1,2-DAG 3-β-galactosyltransferase (or MGDG synthase) that transfers galactose from UDP-Gal to DAG via a β1 → 3 glycosidic linkage [38]. MGDG synthase 1 catalyzes the synthesis of eukaryotic and prokaryotic MGDG molecules in vitro with no apparent specificity for either structure [38] and is at the origin of the majority of the MGDG synthesized in standard

#### **Figure 6.**

*Biosynthesis of glycerolipids according to the prokaryotic pathway (MGDG, DGDG, SQDG, and PG). The enzymes involved are: (1) glycerol-3-phosphate acyl transferase; (2) 1-acyl-glycerol-3-phosphoacyltransferase; (3) phosphatidate phosphatase; (4) MGDG synthase; (5) SQDG synthase; (6) phosphatidate cytidyl transferase; (7) CDP-DAG: glycerol-3-phospho-cytidylyltransferase; (8) phosphatidate glycerophosphatase; d: desaturase.*

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lipids (triacylglycerols TAG).

*Plant Lipid Metabolism*

*4.1.2 Synthesis of the DGDG*

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

condition. In contrast, MGDG synthase 2 and 3 would be localized in the outer membrane [38]. These two enzymes have a better affinity for eukaryotic DAG (C18: 2/ C18: 2) [38] and would likely be in the supply of MGDG for synthesis of DGDG [39].

plastids, presumably in the outer membrane of the envelope [41].

*4.1.3 Synthesis of sulfoquinovosyl-diacylglycerol SQDG*

that transfers SQ from UDP-SQ to a DAG molecule [42].

*4.1.4 Synthesis of phosphatidylglycerol (PG)*

activities have been detected [43].

A small proportion of MGDGs are again glycosylated by DGDG synthase (EC 2.4.1.241) to form DGDG. Two enzymes catalyze DGDG synthesis by adding Gal from UDP-Gal to MGDG via α1 → 6 glycosidic linkage [40]. DGDG synthase1 acts preferentially on MGDG C18/C18, whereas DGDG synthase2 seems to have an affinity for MGDG with C16/18 [41]. These two enzymes would be localized in

Similarly, a sulfolipid synthase (EC 3.13.1.1) catalyzes the attachment of UDPsulfoquinovose (UDP-SQ ) to the *sn*-3 position of DAG to form SQDG. The first step in the synthesis of SQDG or sulfolipid is the formation of UDP-SQ, a polar donor group [32]. The second reaction is catalyzed by a sulfolipid synthase (EC 3.13.1.1)

Phosphatidic acid (PA) is also a substrate for CDP-DAG synthase (EC 2.7.7.41) to form CDP-DAG, the precursor of PG synthesis (**Figure 5**). In chloroplasts, PG is generated in the inner membrane of the envelope where phosphatidylglycerolphosphate synthase and phosphatidylglycerol-phosphate phosphatase (EC 3.1.3.27)

The fatty acids that make up the various glycerolipids formed in the plastid are characterized by a high degree of unsaturations introduced by the various fatty acid desaturases (FAD6, FAD7 and FAD8, EC 1.14.19) to generate polyunsaturated fatty

A major proportion of palmitic and oleic acids are transported as CoA esters outside the chloroplast to be incorporated at the endoplasmic reticulum (ER) into the phospholipids (PC, PE, PI, and PS) (**Figure 6**). ER is the main site for the synthesis of phospholipids and triacylglycerol, which derive from lysophosphatidic

In plant, the glycerophosphate acyltransferase (GPAT) family is involved in the first reaction leading to LPA synthesis of the eukaryotic pathway [35]. In the second reaction, cytosolic lysophosphatidic acid acyl transferase (LPAAT2, EC 2.3.1.23) specifically incorporates oleic acid at the *sn*-2 position of LPA, which is the specific

Most of the flow of chloroplast-exported fatty acids is incorporated in phosphatidylcholine (PC) by a mechanism called "acyl editing" [40]. This mechanism consists of a deacylation-reacylation cycle of the PC which makes it possible to exchange acyls present on the PC with activated FAs taken from a cytosolic pool of free acyl CoA. The oleate exported from plastids, in the form of oleoyl CoA, is used as a substrate for the synthesis of polyunsaturated fatty acids which are inserted either in membrane lipids (PC, PE, and PI) or in storage

acids (PUFA) necessary for the proper functioning of plastids [43].

acid (LPA) as for the prokaryotic pathway (**Figure 7**).

signature glycerolipids from the eukaryotic pathway.

**4.2 Synthesis of glycerophospholipids in the endoplasmic reticulum**

### *Plant Lipid Metabolism DOI: http://dx.doi.org/10.5772/intechopen.81355*

condition. In contrast, MGDG synthase 2 and 3 would be localized in the outer membrane [38]. These two enzymes have a better affinity for eukaryotic DAG (C18: 2/ C18: 2) [38] and would likely be in the supply of MGDG for synthesis of DGDG [39].

## *4.1.2 Synthesis of the DGDG*

*Advances in Lipid Metabolism*

(sulfoquinovosyldiacylglycerol) (**Figure 6**).

ture, SQDG, MGDG, and DGDG (**Figure 5**).

*4.1.1 Synthesis of monogalactosyl diacylglycerol (MGDG)*

a glycerol-3-phosphate at position *sn*-1 by an acyl ACP-glycerol 3 phosphate acyltransferase (EC 2.3.1.1), soluble in the stroma of the plastid [35]. Lysophosphatidic acid (LPA) is thus formed. A second plastid-related plastid acyltransferase, the LPA-ACP acyltransferase, catalyzes the esterification of palmitoyl-ACP at the *sn*-2 position (LPAAT1; EC 2.3.1.51) [36]. This results in the synthesis of 18:1/16:0-PA. Phosphatidic acid (PA) can either be converted to CDP-DAG by the action of a CTP-phosphatidate cytidylyltransferase (EC 2.7.7.41) which catalyzes the reaction between PA and CTP to form CDP-DAG and pyrophosphate or dephosphorylated to diacylglycerol (DAG) by phosphatidate phosphatase (PAP; EC 3.1.3.4). CDP-DAG will be used for the synthesis of phosphatidyl glycerol (PG) of the plastid [35] and DAG can be used for the synthesis of galactolipids (MGDG, DGDG) or a sulfolipid

From the phyllogenetic point of view, the difference between so-called "C16:3" and "C18:3" plants is related to the presence of plastid phosphatidate phosphatase in "C16: 3" plants, lost during evolution in "C18:3" plants. The chloroplast enzyme is clearly different from other phosphatidate phosphatases in the cell because it is membrane-bound, strongly associated with the inner membrane of the envelope, has an optimum alkaline pH, and is inhibited by cations such as Mg2+ [37]. The DAG thus produced (18/16 DAG) is at the origin of the glycolipids of prokaryotic struc-

MGDG is synthesized in a single step by a 1,2-DAG 3-β-galactosyltransferase (or MGDG synthase) that transfers galactose from UDP-Gal to DAG via a β1 → 3 glycosidic linkage [38]. MGDG synthase 1 catalyzes the synthesis of eukaryotic and prokaryotic MGDG molecules in vitro with no apparent specificity for either structure [38] and is at the origin of the majority of the MGDG synthesized in standard

*Biosynthesis of glycerolipids according to the prokaryotic pathway (MGDG, DGDG, SQDG, and PG). The enzymes involved are: (1) glycerol-3-phosphate acyl transferase; (2) 1-acyl-glycerol-3-phosphoacyltransferase; (3) phosphatidate phosphatase; (4) MGDG synthase; (5) SQDG synthase; (6) phosphatidate cytidyl transferase; (7) CDP-DAG: glycerol-3-phospho-cytidylyltransferase; (8) phosphatidate glycerophosphatase; d:* 

**48**

**Figure 6.**

*desaturase.*

A small proportion of MGDGs are again glycosylated by DGDG synthase (EC 2.4.1.241) to form DGDG. Two enzymes catalyze DGDG synthesis by adding Gal from UDP-Gal to MGDG via α1 → 6 glycosidic linkage [40]. DGDG synthase1 acts preferentially on MGDG C18/C18, whereas DGDG synthase2 seems to have an affinity for MGDG with C16/18 [41]. These two enzymes would be localized in plastids, presumably in the outer membrane of the envelope [41].

## *4.1.3 Synthesis of sulfoquinovosyl-diacylglycerol SQDG*

Similarly, a sulfolipid synthase (EC 3.13.1.1) catalyzes the attachment of UDPsulfoquinovose (UDP-SQ ) to the *sn*-3 position of DAG to form SQDG. The first step in the synthesis of SQDG or sulfolipid is the formation of UDP-SQ, a polar donor group [32]. The second reaction is catalyzed by a sulfolipid synthase (EC 3.13.1.1) that transfers SQ from UDP-SQ to a DAG molecule [42].

## *4.1.4 Synthesis of phosphatidylglycerol (PG)*

Phosphatidic acid (PA) is also a substrate for CDP-DAG synthase (EC 2.7.7.41) to form CDP-DAG, the precursor of PG synthesis (**Figure 5**). In chloroplasts, PG is generated in the inner membrane of the envelope where phosphatidylglycerolphosphate synthase and phosphatidylglycerol-phosphate phosphatase (EC 3.1.3.27) activities have been detected [43].

The fatty acids that make up the various glycerolipids formed in the plastid are characterized by a high degree of unsaturations introduced by the various fatty acid desaturases (FAD6, FAD7 and FAD8, EC 1.14.19) to generate polyunsaturated fatty acids (PUFA) necessary for the proper functioning of plastids [43].

## **4.2 Synthesis of glycerophospholipids in the endoplasmic reticulum**

A major proportion of palmitic and oleic acids are transported as CoA esters outside the chloroplast to be incorporated at the endoplasmic reticulum (ER) into the phospholipids (PC, PE, PI, and PS) (**Figure 6**). ER is the main site for the synthesis of phospholipids and triacylglycerol, which derive from lysophosphatidic acid (LPA) as for the prokaryotic pathway (**Figure 7**).

In plant, the glycerophosphate acyltransferase (GPAT) family is involved in the first reaction leading to LPA synthesis of the eukaryotic pathway [35]. In the second reaction, cytosolic lysophosphatidic acid acyl transferase (LPAAT2, EC 2.3.1.23) specifically incorporates oleic acid at the *sn*-2 position of LPA, which is the specific signature glycerolipids from the eukaryotic pathway.

Most of the flow of chloroplast-exported fatty acids is incorporated in phosphatidylcholine (PC) by a mechanism called "acyl editing" [40]. This mechanism consists of a deacylation-reacylation cycle of the PC which makes it possible to exchange acyls present on the PC with activated FAs taken from a cytosolic pool of free acyl CoA. The oleate exported from plastids, in the form of oleoyl CoA, is used as a substrate for the synthesis of polyunsaturated fatty acids which are inserted either in membrane lipids (PC, PE, and PI) or in storage lipids (triacylglycerols TAG).

#### **Figure 7.**

*Biosynthesis of glycerolipids according to the eukaryotic pathway (PC, PE, PI, and PS). The enzymes involved are: (1) glycerol-3-phosphate acyl transferase; (2) 1-acyl-glycerol-3-phosphoacyltransferase; (3) phosphatidate phosphatase; (4) CDP-choline: DAG choline phosphotransferase; (5) CDP-ethanolamine: DAG ethanolamine phosphotransferase or PE synthase; (6) phosphatidate cytidyl transferase; (7) PS synthase; (8) PI synthase; (9) PS decarboxylase; (10) N-methyltransferase.*

In general, the synthesis of phospholipids is separated into three pathways: the phospholipids derived from cytidine diphosphate (CDP)-DAG (PI, PS), those derived from DAG (PC, PE) (**Figure 6**), and those from exchange of polar heads belonging to other phospholipids.

#### *4.2.1 Phospholipids derived from CDP-DAG: PI and PS*

PA can be converted to CDP-DAG by the action of a CTP phosphatidate cytidylyltransferase. This enzyme catalyzes the reaction between a eukaryotic PA molecule and a CTP molecule to form CDP-DAG and pyrophosphate.

Phosphoinositides are an important group of complex structure. PI represents 93% of phosphoinositides, while PIP (mainly PI-3P and PI-4P) and PIP2 (PI-(4,5) P2) represent less than 1%. These phosphoinositides play a major role in signaling processes. PI synthesis is catalyzed by PI synthase from free inositol and CDP-DAG. PI-3P and PI-4P are formed by phosphorylation of PI, respectively, by PI 3- and PI 4-kinases. Finally, PIP2 is formed from PI-4P by PI-4P 5-kinase activity (**Figure 7**).

PS synthase catalyzes the addition of serine to CDP-DAG [44].

#### *4.2.2 Lipids derived from DAG: PE and PC*

The plants synthesize ethanolamine by decarboxylation of serine [45], by serine decarboxylase which is a soluble, plant-specific enzyme. The synthesized free ethanolamine is then phosphorylated by an ethanolamine kinase, specific for ethanolamine different from choline kinase [46]. Phosphoethanolamine is then converted to CDP-ethanolamine by a CTP: phosphoethanolamine cytidyl transferase. The last step of PE synthesis is catalyzed by a CDP-ethanolamine: DAG ethanolamine

**51**

*Plant Lipid Metabolism*

**5. TAG biosynthesis**

pollen grains [49, 50].

can modify the acyl chain.

dent times during angiosperm evolution.

specialized acyl-ACP thioesterase [52].

synthesis outside the plastids [53].

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

synthesizes both PE and PC [47] (**Figure 7**).

way using CDP-choline is preponderant [48].

phosphotransferase. This enzyme is an amino alcohol phosphotransferase that

PC can also be synthesized by two different ways, either by methylation of PE with PE-N-methyltransferase, or by the addition of CDP-choline on DAG. The path-

The synthesis of all these lipids, phospholipids, and glycolipids is localized in specific membranes. However, a large part of the lipids thus generated is present in other membranes than those in which they are synthesized (vacuole, plasma membrane, and thylakoid). The cell therefore has specific lipid transport mechanisms.

TAGs are neutral lipids and are the major component of oilseed oil. These storage lipids represent the main source of carbon and energy mobilized during germination. Other tissues can also accumulate TAGs, such as senescence leaves or

Their biosynthesis occurs at the ER membrane during the storage accumulation phase after embryogenesis. The TAGs result from the esterification at the *sn*-3 position of DAG of fatty acid from the pool of cytosolic acyl CoA by the action of diacylglycerol acyltransferase (DGAT, EC 2.3.1.20) or phospholipid: diacylglycerol acyltransferase (PDAT, EC 2.3.1.158) [17]. The acyl CoAs can be released from PC after desaturation with lyso-PC acyltransferase and reincorporated into the DAG-TAG chain. This allows for a renewal of the fatty acid composition of TAGs [51].

More than 300 different fatty acids are known to occur in seed TAG. Chain length may range from less than 8 to over 22 carbons. The position and number of double bonds may also be unusual, and hydroxy, epoxy, or other functional groups

The synthesis of these unusual fatty acids involves just one additional or alternative enzymatic step from primary lipid metabolism. All the enzymes identified to date that are involved in unusual fatty acid biosynthesis are structurally related to enzymes of primary lipid metabolism. Many of the unusual fatty acids are found in taxonomically dispersed families implying that the recruitment of enzymes for the synthesis of these unusual fatty acids might have occurred a number of indepen-

Plants that synthesize **unusual monounsaturated fatty acids** have an additional desaturase, which is closely related to the Δ9-desaturase but introduces a double bond at a different location on the acyl-ACP. In coriander (*Coriandrum sativum*), petroselinic acid is synthesized by a desaturase that introduces a double bond between carbons 4 and 5 of a C16 acyl-ACP (Δ4-desaturase). This fatty acid is then extended by two carbons and cleaved from ACP to produce the free fatty acid. These last two steps are thought to require a specialized condensing enzyme and a

Plants that synthesize **medium-chain fatty** acids have several thioesterases. Indeed, plants that produce seeds with high concentrations of 8 to 14 carbon atoms, like *Cuphea lanceolata* rich in decanoic acid (C10: 0) *Umbellularia californica* rich in laurate (C12: 0) contain specific thioesterase for medium fatty acid chains. By removing acyl groups from ACP prematurely, the medium-chain thioesterases simultaneously prevent their further elongation and release them for triacylglycerol

**5.1 Seed triacylglycerols often contain unusual fatty acids**

phosphotransferase. This enzyme is an amino alcohol phosphotransferase that synthesizes both PE and PC [47] (**Figure 7**).

PC can also be synthesized by two different ways, either by methylation of PE with PE-N-methyltransferase, or by the addition of CDP-choline on DAG. The pathway using CDP-choline is preponderant [48].

The synthesis of all these lipids, phospholipids, and glycolipids is localized in specific membranes. However, a large part of the lipids thus generated is present in other membranes than those in which they are synthesized (vacuole, plasma membrane, and thylakoid). The cell therefore has specific lipid transport mechanisms.

## **5. TAG biosynthesis**

*Advances in Lipid Metabolism*

belonging to other phospholipids.

*(9) PS decarboxylase; (10) N-methyltransferase.*

**Figure 7.**

*4.2.2 Lipids derived from DAG: PE and PC*

*4.2.1 Phospholipids derived from CDP-DAG: PI and PS*

and a CTP molecule to form CDP-DAG and pyrophosphate.

In general, the synthesis of phospholipids is separated into three pathways: the phospholipids derived from cytidine diphosphate (CDP)-DAG (PI, PS), those derived from DAG (PC, PE) (**Figure 6**), and those from exchange of polar heads

*Biosynthesis of glycerolipids according to the eukaryotic pathway (PC, PE, PI, and PS). The enzymes involved are: (1) glycerol-3-phosphate acyl transferase; (2) 1-acyl-glycerol-3-phosphoacyltransferase; (3) phosphatidate phosphatase; (4) CDP-choline: DAG choline phosphotransferase; (5) CDP-ethanolamine: DAG ethanolamine phosphotransferase or PE synthase; (6) phosphatidate cytidyl transferase; (7) PS synthase; (8) PI synthase;* 

PA can be converted to CDP-DAG by the action of a CTP phosphatidate cytidylyltransferase. This enzyme catalyzes the reaction between a eukaryotic PA molecule

Phosphoinositides are an important group of complex structure. PI represents 93% of phosphoinositides, while PIP (mainly PI-3P and PI-4P) and PIP2 (PI-(4,5) P2) represent less than 1%. These phosphoinositides play a major role in signaling processes. PI synthesis is catalyzed by PI synthase from free inositol and CDP-

DAG. PI-3P and PI-4P are formed by phosphorylation of PI, respectively, by PI 3- and PI 4-kinases. Finally, PIP2 is formed from PI-4P by PI-4P 5-kinase activity (**Figure 7**).

The plants synthesize ethanolamine by decarboxylation of serine [45], by serine decarboxylase which is a soluble, plant-specific enzyme. The synthesized free ethanolamine is then phosphorylated by an ethanolamine kinase, specific for ethanolamine different from choline kinase [46]. Phosphoethanolamine is then converted to CDP-ethanolamine by a CTP: phosphoethanolamine cytidyl transferase. The last step of PE synthesis is catalyzed by a CDP-ethanolamine: DAG ethanolamine

PS synthase catalyzes the addition of serine to CDP-DAG [44].

**50**

TAGs are neutral lipids and are the major component of oilseed oil. These storage lipids represent the main source of carbon and energy mobilized during germination. Other tissues can also accumulate TAGs, such as senescence leaves or pollen grains [49, 50].

Their biosynthesis occurs at the ER membrane during the storage accumulation phase after embryogenesis. The TAGs result from the esterification at the *sn*-3 position of DAG of fatty acid from the pool of cytosolic acyl CoA by the action of diacylglycerol acyltransferase (DGAT, EC 2.3.1.20) or phospholipid: diacylglycerol acyltransferase (PDAT, EC 2.3.1.158) [17]. The acyl CoAs can be released from PC after desaturation with lyso-PC acyltransferase and reincorporated into the DAG-TAG chain. This allows for a renewal of the fatty acid composition of TAGs [51].

### **5.1 Seed triacylglycerols often contain unusual fatty acids**

More than 300 different fatty acids are known to occur in seed TAG. Chain length may range from less than 8 to over 22 carbons. The position and number of double bonds may also be unusual, and hydroxy, epoxy, or other functional groups can modify the acyl chain.

The synthesis of these unusual fatty acids involves just one additional or alternative enzymatic step from primary lipid metabolism. All the enzymes identified to date that are involved in unusual fatty acid biosynthesis are structurally related to enzymes of primary lipid metabolism. Many of the unusual fatty acids are found in taxonomically dispersed families implying that the recruitment of enzymes for the synthesis of these unusual fatty acids might have occurred a number of independent times during angiosperm evolution.

Plants that synthesize **unusual monounsaturated fatty acids** have an additional desaturase, which is closely related to the Δ9-desaturase but introduces a double bond at a different location on the acyl-ACP. In coriander (*Coriandrum sativum*), petroselinic acid is synthesized by a desaturase that introduces a double bond between carbons 4 and 5 of a C16 acyl-ACP (Δ4-desaturase). This fatty acid is then extended by two carbons and cleaved from ACP to produce the free fatty acid. These last two steps are thought to require a specialized condensing enzyme and a specialized acyl-ACP thioesterase [52].

Plants that synthesize **medium-chain fatty** acids have several thioesterases. Indeed, plants that produce seeds with high concentrations of 8 to 14 carbon atoms, like *Cuphea lanceolata* rich in decanoic acid (C10: 0) *Umbellularia californica* rich in laurate (C12: 0) contain specific thioesterase for medium fatty acid chains. By removing acyl groups from ACP prematurely, the medium-chain thioesterases simultaneously prevent their further elongation and release them for triacylglycerol synthesis outside the plastids [53].

Seeds of *Ricinus communis* L. are the source of castor oil, used for the production of high-quality lubricants due to its high proportion of the unusual fatty acid ricinoleic acid. Castor bean seed oil contains 90% of the unusual hydroxy-fatty acid. Castor bean seeds contain an oleate hydroxylase which is structurally similar to extraplastidial membrane-bound Δ12-desaturases (FAD2), and only four amino acid substitutions are needed to convert an 18:1-desaturase into an 18:1-hydroxylase [54]. The synthesis of these fatty acids is thought to take place on the endoplasmic reticulum and use fatty acids esterified to the major membrane lipid phosphatidylcholine as a substrate.

Borage (*Borago officinalis L*.) seeds and evening primrose (*Oenothera biennis* L.) seeds are rich in γlinolenic acid (Δ6, 9, 12), respectively, from 22 to 25% and from 8 to 10%, an essential fatty acid. Its synthesis takes place in the RE during the formation of the seed. The precursor is a linoleoyl-PC and the desaturation is catalyzed by a D6 desaturase [55].

Very long-chain fatty acids (AGTLCs, containing more than 18 carbons) are used in the biosynthesis of many lipids involved in seed storage and waxes. Very long-chain fatty acids (VLCFAs) are synthesized in the following by-products of elongation of a C18 fatty acyl precursor by two carbons originating from malonyl CoA. Each elongation step requires four enzymatic reactions: condensation between an acyl precursor and malonyl-CoA, followed by a reduction, dehydration, and another reduction.

## **6. Conclusion**

The reason for the great diversity in plant storage oils is unknown. The special physical or chemical properties of the "unusual" plant fatty acids have been exploited for centuries. Many of the unusual fatty acids have properties that are valuable as renewable feedstocks for the chemical industry. Medium fatty acids (lauric acid) are the ingredients of a soap or shampoo. VLCFAs like erucic acid (C22:1) can be used as a lubricant or participate in the formation of plastic film. Hydroxy fatty acids such as ricinoleic acid could be a source of biodiesel.

These unusual fatty acids synthesized by spontaneous plants are therefore obtained in small quantities. In order to obtain these fatty acids regularly and in large quantities for industrial use, it will either be necessary to domesticate the plant or introduce the specific gene of the nonconventional fatty acid into an oleaginous plant grown to obtain sufficient yields for industrial uses.

## **Author details**

## Fatiha AID

Laboratoire de Biologie et Physiologie des Organismes (LBPO), Université des Sciences et de la technologie Houari Boumediene (USTHB), Alger, Algerie

\*Address all correspondence to: faid@usthb.dz

© 2019 The Author(s). Licensee IntechOpen. 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.

**53**

*Plant Lipid Metabolism*

**References**

2007

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

[1] Gunstone FD, Harwood JL, Dijkstra AJ, editors. The Lipid Handbook. 3rd ed. Boca Raton, Florida: Taylor & Francis;

[9] Lin M, Oliver DJ. The role of acetylcoenzyme A synthetase in *Arabidopsis*. Plant Physiology. 2008;**147**:1822-1829

[10] Li-Beisson Y, Shorrosh B, Beisson F, Andersson MX, Arondel V, Bates PD, et al. Acyl-lipid metabolism. Arabidopsis eBook. 2010;**11**:01-65

[11] Roughan PG, Ohlrogge JB. Evidence that isolated chloroplasts contain an integrated lipid-synthesizing assembly that channels acetate into long-chain fatty acids. Plant Physiology.

[12] Sasaki Y, Nagano Y. Plant acetylCoA carboxylase: Structure, biosynthesis, regulation and gene manipulation for plant breeding. Bioscience, Biotechnology, and Biochemistry.

[13] Konishi T, Shinohara K, Yamada K, Sasaki Y. Acetyl-CoA carboxylase in higher plants: Most plants other than *Gramineae* have both the prokaryotic and the eukaryotic form of this enzyme. Plant and Cell Physiology.

Trémolières A, Aïd F, Benhassaine-Kesri G. Sethoxydim affects lipid synthesis and acetyl-CoA carboxylase activity in soybean. Journal of Experimental

[15] Harwood JL. Fatty acid biosynthesis. In: Murphy DJ, editor. Plant Lipids: Biology, Utilisation and Manipulation. Oxford: Blackwell Publishing; 2005.

[16] Harwood JL. Recent advances in the biosynthesis of plant fatty acids. Biochimica et Biophysica Acta.

[17] Zhang Y, Maximova SN, Guiltinan MJ. Characterization of a

1996;**110**:1239-1247

2004;**68**(6):1175-1184

1996;**37**:117-122

pp. 27-66

1996;**1301**:7-56

[14] Belkebir A, De Paepe R,

Botany. 2006;**57**:3553-3562

[2] Podkowinski J, Jelenska J, Sirikhachornkit A, Zuther E, Haselkorn R, Gornicki P. Expression of cytosolic and plastid acetylcoenzyme A carboxylase genes in young wheat plants. Plant Physiology.

[3] Stumpf PK. The biosynthesis of saturated fatty acids. In: Stumpf PK, Cohn EE, editors. The Biochemistry of Plants. Vol. 9. New York: Acad. Press;

[4] Harwood JL. Plant acyl lipids: Structure, distribution and analysis. In: Stumpf PK, Conn EE, editors. The Biochemistry of Plants. Vol. 4. New York: Academic Press; 1980.

[5] Dubacq JP, Trémolières A. Occurrence and function of

[6] Dubacq JP, Drapier D, Trémolières A. Polyunsaturated fatty acid synthesis by a mixture of chloroplasts and microsomes from spinach leaves: Evidence of two distinct pathways of biosynthesis of trienoic acids. Plant and Cell Physiology.

[7] Kinney AJ. Perspectives on the production of industrial oils in genetically engineered oilseeds. In: Kuo TM, Gardner HW, editors. Lipid Biotechnology. New York: Marcel

Dekker; 2001. pp. 85-93

DOI: 10.1105/tpc.7.7.957

[8] Ohlrogge J, Browse J. Lipid

biosynthesis. Plant Cell. 1995;**7**:957-970.

phosphatidylglycerol containing 3 *trans* hexadecenoic acid in photosynthetic lamellae. Physiologie végétale.

2003;**131**:763-772

1987. pp. 121-136

pp. 1-55

1983;**21**:293-312

1983;**24**:1-9

## **References**

*Advances in Lipid Metabolism*

a D6 desaturase [55].

**6. Conclusion**

**52**

**Author details**

Fatiha AID

provided the original work is properly cited.

\*Address all correspondence to: faid@usthb.dz

© 2019 The Author(s). Licensee IntechOpen. 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,

Laboratoire de Biologie et Physiologie des Organismes (LBPO), Université des Sciences et de la technologie Houari Boumediene (USTHB), Alger, Algerie

Seeds of *Ricinus communis* L. are the source of castor oil, used for the production of high-quality lubricants due to its high proportion of the unusual fatty acid ricinoleic acid. Castor bean seed oil contains 90% of the unusual hydroxy-fatty acid. Castor bean seeds contain an oleate hydroxylase which is structurally similar to extraplastidial membrane-bound Δ12-desaturases (FAD2), and only four amino acid substitutions are needed to convert an 18:1-desaturase into an 18:1-hydroxylase [54]. The synthesis of these fatty acids is thought to take place on the endoplasmic reticulum and use fatty acids esterified to the major membrane lipid phosphatidylcholine as a substrate.

Borage (*Borago officinalis L*.) seeds and evening primrose (*Oenothera biennis* L.) seeds are rich in γlinolenic acid (Δ6, 9, 12), respectively, from 22 to 25% and from 8 to 10%, an essential fatty acid. Its synthesis takes place in the RE during the formation of the seed. The precursor is a linoleoyl-PC and the desaturation is catalyzed by

Very long-chain fatty acids (AGTLCs, containing more than 18 carbons) are used in the biosynthesis of many lipids involved in seed storage and waxes. Very long-chain fatty acids (VLCFAs) are synthesized in the following by-products of elongation of a C18 fatty acyl precursor by two carbons originating from malonyl CoA. Each elongation step requires four enzymatic reactions: condensation between an acyl precursor and malonyl-CoA, followed by a reduction, dehydration, and another reduction.

The reason for the great diversity in plant storage oils is unknown. The special physical or chemical properties of the "unusual" plant fatty acids have been exploited for centuries. Many of the unusual fatty acids have properties that are valuable as renewable feedstocks for the chemical industry. Medium fatty acids (lauric acid) are the ingredients of a soap or shampoo. VLCFAs like erucic acid (C22:1) can be used as a lubricant or participate in the formation of plastic film.

These unusual fatty acids synthesized by spontaneous plants are therefore obtained in small quantities. In order to obtain these fatty acids regularly and in large quantities for industrial use, it will either be necessary to domesticate the plant or introduce the specific gene of the nonconventional fatty acid into an oleaginous

Hydroxy fatty acids such as ricinoleic acid could be a source of biodiesel.

plant grown to obtain sufficient yields for industrial uses.

[1] Gunstone FD, Harwood JL, Dijkstra AJ, editors. The Lipid Handbook. 3rd ed. Boca Raton, Florida: Taylor & Francis; 2007

[2] Podkowinski J, Jelenska J, Sirikhachornkit A, Zuther E, Haselkorn R, Gornicki P. Expression of cytosolic and plastid acetylcoenzyme A carboxylase genes in young wheat plants. Plant Physiology. 2003;**131**:763-772

[3] Stumpf PK. The biosynthesis of saturated fatty acids. In: Stumpf PK, Cohn EE, editors. The Biochemistry of Plants. Vol. 9. New York: Acad. Press; 1987. pp. 121-136

[4] Harwood JL. Plant acyl lipids: Structure, distribution and analysis. In: Stumpf PK, Conn EE, editors. The Biochemistry of Plants. Vol. 4. New York: Academic Press; 1980. pp. 1-55

[5] Dubacq JP, Trémolières A. Occurrence and function of phosphatidylglycerol containing 3 *trans* hexadecenoic acid in photosynthetic lamellae. Physiologie végétale. 1983;**21**:293-312

[6] Dubacq JP, Drapier D, Trémolières A. Polyunsaturated fatty acid synthesis by a mixture of chloroplasts and microsomes from spinach leaves: Evidence of two distinct pathways of biosynthesis of trienoic acids. Plant and Cell Physiology. 1983;**24**:1-9

[7] Kinney AJ. Perspectives on the production of industrial oils in genetically engineered oilseeds. In: Kuo TM, Gardner HW, editors. Lipid Biotechnology. New York: Marcel Dekker; 2001. pp. 85-93

[8] Ohlrogge J, Browse J. Lipid biosynthesis. Plant Cell. 1995;**7**:957-970. DOI: 10.1105/tpc.7.7.957

[9] Lin M, Oliver DJ. The role of acetylcoenzyme A synthetase in *Arabidopsis*. Plant Physiology. 2008;**147**:1822-1829

[10] Li-Beisson Y, Shorrosh B, Beisson F, Andersson MX, Arondel V, Bates PD, et al. Acyl-lipid metabolism. Arabidopsis eBook. 2010;**11**:01-65

[11] Roughan PG, Ohlrogge JB. Evidence that isolated chloroplasts contain an integrated lipid-synthesizing assembly that channels acetate into long-chain fatty acids. Plant Physiology. 1996;**110**:1239-1247

[12] Sasaki Y, Nagano Y. Plant acetylCoA carboxylase: Structure, biosynthesis, regulation and gene manipulation for plant breeding. Bioscience, Biotechnology, and Biochemistry. 2004;**68**(6):1175-1184

[13] Konishi T, Shinohara K, Yamada K, Sasaki Y. Acetyl-CoA carboxylase in higher plants: Most plants other than *Gramineae* have both the prokaryotic and the eukaryotic form of this enzyme. Plant and Cell Physiology. 1996;**37**:117-122

[14] Belkebir A, De Paepe R, Trémolières A, Aïd F, Benhassaine-Kesri G. Sethoxydim affects lipid synthesis and acetyl-CoA carboxylase activity in soybean. Journal of Experimental Botany. 2006;**57**:3553-3562

[15] Harwood JL. Fatty acid biosynthesis. In: Murphy DJ, editor. Plant Lipids: Biology, Utilisation and Manipulation. Oxford: Blackwell Publishing; 2005. pp. 27-66

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[18] Fox BG, Shanklin J, Somerville C, Munck E. Stearoylacyl carrier protein D9 desaturase from *Ricinus communis* is a di-iron-oxo protein. Proceedings of the National Academy of Sciences of the United States of America. 1993;**90**:2486-2490

[19] Shanklin J, Somerville C. Stearoylacyl-carrier-protein desaturase from higher plants is structurally unrelated to the animal and fungal homologs (fatty acid desaturation/cDNA clone/ lipid unsaturation/fatty acid synthase). Proceedings of the National Academy of Sciences of the United States of America. 1991;**88**:2510-2514

[20] Roughan PG, Slack RC. Cellular organization of glycerolipid metabolism. Annual Review of Plant Physiology. 1982;**33**:97-132

[21] Heemskerk JWM, Wintermans JFGM. Role of the chloroplast in the leaf acid lipid synthesis. Physiologia Plantarum. 1987;**70**:558-568

[22] Löhden I, Frentzen M. Role of plastidial acyl-acyl carrier protein: Glycerol 3 phosphate acyltransferase and acyl-acyl carrier protein hydrolase in channeling the acyl flux through the prokaryotic and eukaryotic pathway. Planta. 1988;**176**:506-5012

[23] Sato N, Murata N. Lipid biosynthesis in blue green algae Anabaena variabilis. II Fatty acids and molecular species. Biochimica et Biophysica Acta. 1982;**710**:279-289

[24] Mazliak P, Justin AM, Demandre C, Chicha A. Specificity of some enzymes involved in glycerolipid biosynthesis. In: Cherif A et al., editors. Metabolism, Structure and Utilization

of Plant Lipids. Tunis: Centre National Pedagogique; 1992. pp. 3-17

[25] Heemskerck JWM, Schmidt H, Hammer V, Wintermans JFGM. Biosynthesis and desaturation of prokaryotic galactolipids in and isolated chloroplasts from spinach leaves. Plant Physiology. 1991;**96**:144-152

[26] Ohlogge JB, Browse J, Somerville CR. The genetic of plant lipids. Biochimica et Biophysica Acta. 1991;**1082**:1-26

[27] Block MA, Douce R, Joyard J, Rolland N. Chloroplast envelope membranes: A dynamic interface between plastids and the cytosol. Photosynthesis Research. 2007;**92**:225-244

[28] Trémolières A, Dubacq JP, Drapier D, Muller M, Mazliak P. In vitro cooperation between plastids and microsomes in the leaf lipids. FEBS Letters. 1980;**114**:135-138

[29] Bates PD, Fatihi A, Snapp AR, Carlsson AS, Browse JA, Lu C. Acyl editing and headgroup exchange are the major mechanisms that direct polyunsaturated fatty acid flux into triacylglycerols. Plant Physiology. 2012;**160**:1530-1539

[30] Serghini-Caid H, Demandre C, Justin AM, Mazliak P. Oleoyl-PCmolecular species desaturated in pea leaf microsomes. Possible substrates of oleate desaturase in other green leaves. Plant Science. 1988;**54**:93-101

[31] Benning C. Mechanisms of lipid transport involved in organelle biogenesis in plant cells. Annual Review of Cell and Developmental Biology. 2009;**25**:71-91

[32] Wallis JG, Browse J. Mutants of *Arabidopsis* reveal many roles for membrane lipids. Progress in Lipid Research. 2002;**41**:254-278

**55**

*Plant Lipid Metabolism*

fpls.2013.00469

2006;**47**:296-309

2004;**134**:1206-1216

*DOI: http://dx.doi.org/10.5772/intechopen.81355*

[39] Kelly AA, Froehlich JE, Dörmann P. Disruption of the two digalactosyldiacylglycerol synthase genes DGD1 and DGD2 in *Arabidopsis* reveals the existence of an additional enzyme of galactolipid synthesis. Plant

Cell. 2003;**15**:2694-2706

2001;**276**:3941-3946

2002;**99**:5732-5737

[40] Sanda S, Leustek T, Theisen MJ, Garavito RM, Benning C. Recombinant *Arabidopsis* SQD1 converts UDP-glucose and sulfite to the sulfolipid head group precursor UDP-sulfoquinovose in vitro. The Journal of Biological Chemistry.

[41] Yu B, Xu C, Benning C. *Arabidopsis* disrupted in SQD2 encoding sulfolipid synthase is impaired in phosphatelimited growth. Proceedings of the National Academy of Sciences of the United States of America.

[42] Gidda SK, Shockey JM, Rothstein SJ,

localized to the ER and possess distinct

[43] Xu C, Härtel H, Wada H, Hagio M,

[44] Delhaize E, Hebb DM, Richards KD, Lin JM, Ryan PR, Gardner RC. Cloning and expression of a wheat (*Triticum aestivum* L.) phosphatidylserine synthase cDNA. Overexpression in plants alters the composition of phospholipids. The Journal of Biological

Dyer JM, Mullen RT. *Arabidopsis thaliana* GPAT8 and GPAT9 are

ER retrieval signals: Functional divergence of the dilysine ER retrieval motif in plant cells. Plant Physiology and Biochemistry. 2009;**47**:867-879

Yu B, Eakin C, et al. The pgp1 mutant locus of *Arabidopsis* encodes a phosphatidylglycerolphosphate synthase with impaired activity. Plant

Physiology. 2002;**129**:594-604

Chemistry. 1999;**274**:7082-7088

[45] Rontein D, Rhodes D, Hanson AD. Evidence from engineering that decarboxylation of free serine is the major source of ethanolamine moieties

[34] Xu C, Yu B, Cornish AJ, Froehlich JE, Benning C. Phosphatidylglycerol biosynthesis in chloroplasts of *Arabidopsis* mutants deficient in acyl-ACP glycerol-3phosphate acyltransferase. The Plant Journal.

[35] Kim HU, Huang AHC. Plastid lysophosphatidyl acyltransferase is essential for embryo development in *Arabidopsis*. Plant Molecular Biology.

[36] Block MA, Dorne AJ, Joyard J,

[37] Awai K, Maréchal E, Block MA, Brun D, Masuda T, Shimada H, et al. Two types of MGDG synthase genes, found widely in both 16:3 and 18:3 plants, differentially mediate galactolipid syntheses in photosynthetic and nonphotosynthetic tissues in *Arabidopsis thaliana*. Proceedings of the National Academy of Sciences of the United States of America.

[38] Kobayashi K, Awai K, Takamiya K,

characterization of membrane fractions enriched in outer and inner envelope membranes from spinach chloroplasts. II. Biochemical characterization. The Journal of Biological Chemistry.

Douce R. Preparation and

1983;**258**:13281-13286

2001;**98**:10960-10965

2004;**134**:640-648

Ohta H. *Arabidopsis* type B monogalactosyldiacylglycerol synthase genes are expressed during pollen tube growth and induced by phosphate starvation. Plant Physiology.

[33] Shimojima M, Watanabe T, Madoka Y, Koizumi R, Yamamoto MP, Masuda K. Differential regulation of two types of monogalactosyldiacylglycerol synthase in membrane lipid remodeling under phosphate-limited conditions in sesame plants. Frontiers in Plant Science. 2013;**4**:469. DOI: 10.3389/

*Plant Lipid Metabolism DOI: http://dx.doi.org/10.5772/intechopen.81355*

*Advances in Lipid Metabolism*

fpls.2015.00239

1993;**90**:2486-2490

stearoyl-acyl carrier protein desaturase gene family from chocolate tree, *Theobroma cacao* L. Frontiers in Plant Science. 2015;**6**:239. DOI: 10.3389/

of Plant Lipids. Tunis: Centre National

[25] Heemskerck JWM, Schmidt H, Hammer V, Wintermans JFGM. Biosynthesis and desaturation of prokaryotic galactolipids in and isolated chloroplasts from spinach leaves. Plant

Somerville CR. The genetic of plant lipids. Biochimica et Biophysica Acta.

[27] Block MA, Douce R, Joyard J, Rolland N. Chloroplast envelope membranes: A dynamic interface between plastids and the cytosol.

Pedagogique; 1992. pp. 3-17

Physiology. 1991;**96**:144-152

[26] Ohlogge JB, Browse J,

Photosynthesis Research.

Letters. 1980;**114**:135-138

2012;**160**:1530-1539

2009;**25**:71-91

[28] Trémolières A, Dubacq JP, Drapier D, Muller M, Mazliak P. In vitro cooperation between plastids and microsomes in the leaf lipids. FEBS

[29] Bates PD, Fatihi A, Snapp AR, Carlsson AS, Browse JA, Lu C. Acyl editing and headgroup exchange are the major mechanisms that direct polyunsaturated fatty acid flux into triacylglycerols. Plant Physiology.

[30] Serghini-Caid H, Demandre C, Justin AM, Mazliak P. Oleoyl-

[31] Benning C. Mechanisms of lipid transport involved in organelle

[32] Wallis JG, Browse J. Mutants of *Arabidopsis* reveal many roles for membrane lipids. Progress in Lipid

Research. 2002;**41**:254-278

biogenesis in plant cells. Annual Review of Cell and Developmental Biology.

Plant Science. 1988;**54**:93-101

PCmolecular species desaturated in pea leaf microsomes. Possible substrates of oleate desaturase in other green leaves.

1991;**1082**:1-26

2007;**92**:225-244

[18] Fox BG, Shanklin J, Somerville C, Munck E. Stearoylacyl carrier protein D9 desaturase from *Ricinus communis* is a di-iron-oxo protein. Proceedings of the National Academy of Sciences of the United States of America.

[19] Shanklin J, Somerville C. Stearoylacyl-carrier-protein desaturase from higher plants is structurally unrelated to the animal and fungal homologs (fatty acid desaturation/cDNA clone/ lipid unsaturation/fatty acid synthase). Proceedings of the National Academy of Sciences of the United States of America. 1991;**88**:2510-2514

[20] Roughan PG, Slack RC. Cellular

metabolism. Annual Review of Plant

[21] Heemskerk JWM, Wintermans JFGM. Role of the chloroplast in the leaf acid lipid synthesis. Physiologia

[22] Löhden I, Frentzen M. Role of plastidial acyl-acyl carrier protein: Glycerol 3 phosphate acyltransferase and acyl-acyl carrier protein hydrolase in channeling the acyl flux through the prokaryotic and eukaryotic pathway.

organization of glycerolipid

Physiology. 1982;**33**:97-132

Plantarum. 1987;**70**:558-568

Planta. 1988;**176**:506-5012

[23] Sato N, Murata N. Lipid biosynthesis in blue green algae Anabaena variabilis. II Fatty acids and molecular species. Biochimica et Biophysica Acta. 1982;**710**:279-289

Chicha A. Specificity of some enzymes involved in glycerolipid biosynthesis. In: Cherif A et al., editors. Metabolism, Structure and Utilization

[24] Mazliak P, Justin AM, Demandre C,

**54**

[33] Shimojima M, Watanabe T, Madoka Y, Koizumi R, Yamamoto MP, Masuda K. Differential regulation of two types of monogalactosyldiacylglycerol synthase in membrane lipid remodeling under phosphate-limited conditions in sesame plants. Frontiers in Plant Science. 2013;**4**:469. DOI: 10.3389/ fpls.2013.00469

[34] Xu C, Yu B, Cornish AJ, Froehlich JE, Benning C. Phosphatidylglycerol biosynthesis in chloroplasts of *Arabidopsis* mutants deficient in acyl-ACP glycerol-3phosphate acyltransferase. The Plant Journal. 2006;**47**:296-309

[35] Kim HU, Huang AHC. Plastid lysophosphatidyl acyltransferase is essential for embryo development in *Arabidopsis*. Plant Molecular Biology. 2004;**134**:1206-1216

[36] Block MA, Dorne AJ, Joyard J, Douce R. Preparation and characterization of membrane fractions enriched in outer and inner envelope membranes from spinach chloroplasts. II. Biochemical characterization. The Journal of Biological Chemistry. 1983;**258**:13281-13286

[37] Awai K, Maréchal E, Block MA, Brun D, Masuda T, Shimada H, et al. Two types of MGDG synthase genes, found widely in both 16:3 and 18:3 plants, differentially mediate galactolipid syntheses in photosynthetic and nonphotosynthetic tissues in *Arabidopsis thaliana*. Proceedings of the National Academy of Sciences of the United States of America. 2001;**98**:10960-10965

[38] Kobayashi K, Awai K, Takamiya K, Ohta H. *Arabidopsis* type B monogalactosyldiacylglycerol synthase genes are expressed during pollen tube growth and induced by phosphate starvation. Plant Physiology. 2004;**134**:640-648

[39] Kelly AA, Froehlich JE, Dörmann P. Disruption of the two digalactosyldiacylglycerol synthase genes DGD1 and DGD2 in *Arabidopsis* reveals the existence of an additional enzyme of galactolipid synthesis. Plant Cell. 2003;**15**:2694-2706

[40] Sanda S, Leustek T, Theisen MJ, Garavito RM, Benning C. Recombinant *Arabidopsis* SQD1 converts UDP-glucose and sulfite to the sulfolipid head group precursor UDP-sulfoquinovose in vitro. The Journal of Biological Chemistry. 2001;**276**:3941-3946

[41] Yu B, Xu C, Benning C. *Arabidopsis* disrupted in SQD2 encoding sulfolipid synthase is impaired in phosphatelimited growth. Proceedings of the National Academy of Sciences of the United States of America. 2002;**99**:5732-5737

[42] Gidda SK, Shockey JM, Rothstein SJ, Dyer JM, Mullen RT. *Arabidopsis thaliana* GPAT8 and GPAT9 are localized to the ER and possess distinct ER retrieval signals: Functional divergence of the dilysine ER retrieval motif in plant cells. Plant Physiology and Biochemistry. 2009;**47**:867-879

[43] Xu C, Härtel H, Wada H, Hagio M, Yu B, Eakin C, et al. The pgp1 mutant locus of *Arabidopsis* encodes a phosphatidylglycerolphosphate synthase with impaired activity. Plant Physiology. 2002;**129**:594-604

[44] Delhaize E, Hebb DM, Richards KD, Lin JM, Ryan PR, Gardner RC. Cloning and expression of a wheat (*Triticum aestivum* L.) phosphatidylserine synthase cDNA. Overexpression in plants alters the composition of phospholipids. The Journal of Biological Chemistry. 1999;**274**:7082-7088

[45] Rontein D, Rhodes D, Hanson AD. Evidence from engineering that decarboxylation of free serine is the major source of ethanolamine moieties in plants. Plant and Cell Physiology. 2003;**44**:1185-1191

[46] Wharfe J, Harwood JL. Lipid metabolism in germinatingseeds. Purification of ethanolamine kinase from soya bean. Biochimica et Biophysica Acta. 1979;**575**:102-111

[47] Choi YH, Lee JK, Lee CH, Cho SH. cDNA cloning and expression of an aminoalcoholphosphotransferase isoform in Chinese cabbage. Plant and Cell Physiology. 2000;**41**:1080-1084

[48] Tasseva G, Richard L, Zachowski A. Regulation of phosphatidylcholine biosynthesis under saltstress involves choline kinase in *Arabidopsis thaliana*. FEBS Letters. 2004;**566**:115-120

[49] Kaup MT, Froese CD, Thompson JE. A role for diacylglycerol acyltransferase during leaf senescence. Plant Physiology. 2002;**129**:1616-1626

[50] Kim HU, Hsieh K, Ratnayake C, Huang AHC. A novel group of oleosins is present inside the pollen of *Arabidopsis*. The Journal of Biological Chemistry. 2002;**277**:22677-22684

[51] Battey JF, Schmid KM, Ohlrogge JB. Genetic engineering for plant oils: Potential and limitations. Trends in Biotechnology. 1989;**7**:122-125

[52] Grosbois M. Biosynthèse des acides gras au cours du développement du fruit et de la graine du lierre. Phytochemistry. 1971;**10**(6):1261-1273

[53] Pollard MA, Anderson L, Fan C, Hawkins DJ, Davies HM. A specific acyi-ACP thioesterase implicated in laurate production in immature cotyledons of *Umbellularia californica*. In: Quinn PJ, Harwood JL, editors. Plant Lipid Biochemistry, Structure and Utilization. London: Portland; 1990. pp. 163-165

[54] Van de Loo FJ, Turner PBS, Somerville C. An oleate 12 hydroxylase from *Ricinus communis* L. is a fatty acyl desaturase homolog (ricinoleic acid/castor/FAH12/transgenic plants). Proceedings of the National Academy of Sciences of the United States of America. 1995, 1995;**92**:6743-6747

[55] Galle AM, Demandre C, Guerche P, Joseph M, Dubacq JP, Mazliak P, et al. γlinolenic acid biosynthesis in microsomal membranes of developing borage officinal is seeds. In: Chérif A et al., editors. Metabolism, Structure and Utilization of Plant Lipids. Tunis: Centre National Pédagogique; 1992. pp. 185-188

**57**

**Chapter 4**

**Abstract**

compounds.

ascorbate, glutathione

**1. Introduction**

*Shahla Hashemi*

Effect of Nanoparticles on Lipid

The size of the nanoparticles is between 1 and 100 nm. Nanoparticles are widely used in consumer and medical products, as well as in agricultural and industrial applications. The excessive use nanoparticles increases its release into the environment. Plants are an important part of the environment that is affected by nanoparticles. Studies have examined the effect of nanoparticles on plants. The results showed that high concentrations of nanoparticles showed a negative effect. Reactive oxygen species generation is a toxicological mechanism of nanoparticles in plants. When the production of radicals is greater than its removal, oxidative stress occurs. The key indicator of oxidative stress is lipid peroxidation. The unsaturated fatty acids in the cell membrane are a major target for radicals. Radical absorbs hydrogen from unsaturated fatty acids to form water. Therefore, the fatty acid has a non-coupled electron, which is then able to capture oxygen and form a peroxyl radical. Lipid peroxyl radical can lead to a chain of radical production. Enzymatic and nonenzymatic systems exist for the removal of radicals in plants. Enzymatic systems include catalase, guaiacol peroxidase, ascorbate peroxidase, superoxide dismutase, glutathione reductase, and dehydroascorbate reductase. Nonenzymatic systems include ascorbate and carotenoids, glutathione, tocopherol, and phenolic

**Keywords:** nanoparticles, reactive oxygen species, malondialdehyde, catalase,

The nanoparticles have a size of less than 100 nm in at least one dimension. Due to the specific properties of nanoparticles, in particular the high-surfaceto-volume ratio, they have been used for several applications. For example, nanoparticles are used in the fields of biosensors and electronics, cosmetic industries, wastewater treatment, biomedicines, cancer therapy, and targeted drug delivery [1, 2]. The excessive use of nanoparticles results in the release of these materials into the environment. The environment includes plants, the main producers of the food chain, which are affected by nanoparticles. Nanoparticles are absorbed by plants and transmitted to various parts of the plants and affect them. Several factors such as physicochemical properties of nanoparticles, plant species, and exposure conditions contribute to the absorption and transfer of nanoparticles. Size, magnetic properties, surface charge, composition, crystalline state, and surface functionalization are some of the physical properties of nanoparticles that are important in their absorption

Peroxidation in Plants

## **Chapter 4**

*Advances in Lipid Metabolism*

2003;**44**:1185-1191

in plants. Plant and Cell Physiology.

from *Ricinus communis* L. is a fatty acyl desaturase homolog (ricinoleic acid/castor/FAH12/transgenic plants). Proceedings of the National Academy of Sciences of the United States of America. 1995, 1995;**92**:6743-6747

[55] Galle AM, Demandre C, Guerche P, Joseph M, Dubacq JP, Mazliak P, et al. γlinolenic acid biosynthesis in microsomal membranes of developing borage officinal is seeds. In: Chérif A et al., editors. Metabolism, Structure and Utilization of Plant Lipids. Tunis: Centre National Pédagogique; 1992.

pp. 185-188

[46] Wharfe J, Harwood JL. Lipid metabolism in germinatingseeds. Purification of ethanolamine kinase from soya bean. Biochimica et Biophysica Acta. 1979;**575**:102-111

[47] Choi YH, Lee JK, Lee CH, Cho SH. cDNA cloning and expression of an aminoalcoholphosphotransferase isoform in Chinese cabbage. Plant and Cell Physiology. 2000;**41**:1080-1084

[48] Tasseva G, Richard L, Zachowski A. Regulation of phosphatidylcholine biosynthesis under saltstress involves choline kinase in *Arabidopsis thaliana*. FEBS Letters. 2004;**566**:115-120

[49] Kaup MT, Froese CD, Thompson JE. A role for diacylglycerol acyltransferase

[50] Kim HU, Hsieh K, Ratnayake C, Huang AHC. A novel group of oleosins is present inside the pollen of *Arabidopsis*. The Journal of Biological Chemistry. 2002;**277**:22677-22684

[51] Battey JF, Schmid KM, Ohlrogge JB. Genetic engineering for plant oils: Potential and limitations. Trends in Biotechnology. 1989;**7**:122-125

[52] Grosbois M. Biosynthèse des acides gras au cours du développement du fruit et de la graine du lierre. Phytochemistry.

[53] Pollard MA, Anderson L, Fan C, Hawkins DJ, Davies HM. A specific acyi-ACP thioesterase implicated in laurate production in immature cotyledons of *Umbellularia californica*. In: Quinn PJ, Harwood JL, editors. Plant Lipid Biochemistry, Structure and Utilization. London: Portland; 1990. pp. 163-165

[54] Van de Loo FJ, Turner PBS,

Somerville C. An oleate 12 hydroxylase

1971;**10**(6):1261-1273

during leaf senescence. Plant Physiology. 2002;**129**:1616-1626

**56**
