62
should be stored at −80 °C until analysis. Cell disruption (homogenization by mortar pestle or automated techniques) is also important to separate the biomass from
the extracellular matrix of intracellular metabolites and to make them accessible
for different solvents for lipid extraction. After homogenization lipids can be
extracted with appropriate solvent(s) to extract a wide range of lipid metabolites
exhibiting the utmost structural diversity as possible. The liquid-liquid extraction,
organic solvent precipitation, and solid-phase extraction have been used for lipid
extraction in seaweeds, recently reviewed by Maciel et al. (2016). The conventional organic solvent extraction methods based on chloroform/methanol/water,
both Bligh and Dyer (chloroform/methanol; 1/2, v/v) (Bligh and Dyer, 1959) and
Folch method (chloroform/methanol; 2/1, v/v) (Folch et al. 1957), have been invariably used as standard methods for lipid extraction from seaweeds (Galloway et al.
2012; Kumari et al. 2010, 2011, 2013a, b). Other solvents such as dichloromethane
methanol (Graeve et al. 2002), n-butanol (Kim et al. 2007), and diethyl ether
(El-Shoubaky et al. 2008) have also been used but were not so effective. Kumari
et al. (2011) reported that the solvent system comprising of chloroform/methanol/
phosphate buffer is a better combination for extracting lipids because chloroform
and methanol in combination exhibit strong dissolving power for the entire range
of polarity found in lipids, as well as the ability to break up membrane and denature
(-lipo)proteins (Schreiner 2006), and the addition of buffer helped to overcome the
ionic adsorption effects of salt that hinder lipid extraction from seaweeds. An
extraction procedure using methanol/methyl-tert-butyl-ether (MTBE)/water has
also been successfully employed for polar lipidomics of seaweeds (Matyash et al.
2008; Melo et al. 2015). The advantage of this procedure is that MTBE is nontoxic
and noncarcinogenic, so a green solvent for lipid extraction and the lipid-containing phase forms the upper layer during phase separation. Some novel green techniques have also been developed such as supercritical fluid extraction,
microwave-assisted extraction, ultrasound- assisted extraction, and pressurized solvent extraction pulsed electric field-assisted extraction and enzyme-assisted extraction, mostly being tested in microalgae reviewed by Kumari et al. (2013a) and
Maciel et al. (2016) and references therein. The expanding lipidomic field will
surely develop a more suitable environment-friendly, cost-effective, and reproducible green lipid extraction procedures.
Further, it should be taken into account that complete recovery of every lipid
class is difficult to achieve with any known method of lipid extraction. Any incomplete recovery may lead to an incomplete measurement of lipid content in a sample and inconsistencies in the results between inter-laboratory experiments, if
analytical methods are based on external calibration. It is advisable to always add
internal standards (one or more stable isotope labeled compounds) of each lipid
type/class during the extraction procedure, for reliable, accurate, and reproducible
quantification of lipid class of interest (Vaz et al. 2015; Wang et al. 2016). At last,
there are a few precautions to be taken while preparing lipid extracts for lipidomic
analysis. Care should be taken during sample preparation and storage of lipid
extracts from being chemically or enzymatically modified as it negatively influP. Kumari
should be stored at −80 °C until analysis. Cell disruption (homogenization by mortar pestle or automated techniques) is also important to separate the biomass from
the extracellular matrix of intracellular metabolites and to make them accessible
for different solvents for lipid extraction. After homogenization lipids can be
extracted with appropriate solvent(s) to extract a wide range of lipid metabolites
exhibiting the utmost structural diversity as possible. The liquid-liquid extraction,
organic solvent precipitation, and solid-phase extraction have been used for lipid
extraction in seaweeds, recently reviewed by Maciel et al. (2016). The conventional organic solvent extraction methods based on chloroform/methanol/water,
both Bligh and Dyer (chloroform/methanol; 1/2, v/v) (Bligh and Dyer, 1959) and
Folch method (chloroform/methanol; 2/1, v/v) (Folch et al. 1957), have been invariably used as standard methods for lipid extraction from seaweeds (Galloway et al.
2012; Kumari et al. 2010, 2011, 2013a, b). Other solvents such as dichloromethane
methanol (Graeve et al. 2002), n-butanol (Kim et al. 2007), and diethyl ether
(El-Shoubaky et al. 2008) have also been used but were not so effective. Kumari
et al. (2011) reported that the solvent system comprising of chloroform/methanol/
phosphate buffer is a better combination for extracting lipids because chloroform
and methanol in combination exhibit strong dissolving power for the entire range
of polarity found in lipids, as well as the ability to break up membrane and denature
(-lipo)proteins (Schreiner 2006), and the addition of buffer helped to overcome the
ionic adsorption effects of salt that hinder lipid extraction from seaweeds. An
extraction procedure using methanol/methyl-tert-butyl-ether (MTBE)/water has
also been successfully employed for polar lipidomics of seaweeds (Matyash et al.
2008; Melo et al. 2015). The advantage of this procedure is that MTBE is nontoxic
and noncarcinogenic, so a green solvent for lipid extraction and the lipid-containing phase forms the upper layer during phase separation. Some novel green techniques have also been developed such as supercritical fluid extraction,
microwave-assisted extraction, ultrasound- assisted extraction, and pressurized solvent extraction pulsed electric field-assisted extraction and enzyme-assisted extraction, mostly being tested in microalgae reviewed by Kumari et al. (2013a) and
Maciel et al. (2016) and references therein. The expanding lipidomic field will
surely develop a more suitable environment-friendly, cost-effective, and reproducible green lipid extraction procedures.
Further, it should be taken into account that complete recovery of every lipid
class is difficult to achieve with any known method of lipid extraction. Any incomplete recovery may lead to an incomplete measurement of lipid content in a sample and inconsistencies in the results between inter-laboratory experiments, if
analytical methods are based on external calibration. It is advisable to always add
internal standards (one or more stable isotope labeled compounds) of each lipid
type/class during the extraction procedure, for reliable, accurate, and reproducible
quantification of lipid class of interest (Vaz et al. 2015; Wang et al. 2016). At last,
there are a few precautions to be taken while preparing lipid extracts for lipidomic
analysis. Care should be taken during sample preparation and storage of lipid
extracts from being chemically or enzymatically modified as it negatively influP. Kumari