Characteristics of temperature tolerance and ethanol for major industrial strains worldwide used for the production of bioethanol.
\r\n\tThis book will be a self-contained collection of scholarly papers targeting an audience of practicing researchers, academics, PhD students and other scientists. The contents of the book will be written by multiple authors and edited by experts in the field. The area of interest and scope of the project can be described with (but are not limited to) the following keywords: Alcoholism, Depression, Addiction, Blackouts, Relapse, Binge Drinking, Genetic basis, Neurological Aspects, Treatment, Organ Damage.
\r\n\r\n\tAuthors are not limited in terms of topic, but encouraged to present a chapter proposal that best suits their current research efforts. Later, when all chapter proposals are collected, the editor will provide a more specific direction of the book.
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Venkateswarlu",coverURL:"https://cdn.intechopen.com/books/images_new/371.jpg",editedByType:"Edited by",editors:[{id:"58592",title:"Dr.",name:"Arun",surname:"Shanker",slug:"arun-shanker",fullName:"Arun Shanker"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"878",title:"Phytochemicals",subtitle:"A Global Perspective of Their Role in Nutrition and Health",isOpenForSubmission:!1,hash:"ec77671f63975ef2d16192897deb6835",slug:"phytochemicals-a-global-perspective-of-their-role-in-nutrition-and-health",bookSignature:"Venketeshwer Rao",coverURL:"https://cdn.intechopen.com/books/images_new/878.jpg",editedByType:"Edited by",editors:[{id:"82663",title:"Dr.",name:"Venketeshwer",surname:"Rao",slug:"venketeshwer-rao",fullName:"Venketeshwer Rao"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"4816",title:"Face Recognition",subtitle:null,isOpenForSubmission:!1,hash:"146063b5359146b7718ea86bad47c8eb",slug:"face_recognition",bookSignature:"Kresimir Delac and Mislav Grgic",coverURL:"https://cdn.intechopen.com/books/images_new/4816.jpg",editedByType:"Edited by",editors:[{id:"528",title:"Dr.",name:"Kresimir",surname:"Delac",slug:"kresimir-delac",fullName:"Kresimir Delac"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}}]},chapter:{item:{type:"chapter",id:"56367",title:"Phenotypic Characterization of Yeasts Aiming at Bioethanol Production",doi:"10.5772/intechopen.70042",slug:"phenotypic-characterization-of-yeasts-aiming-at-bioethanol-production",body:'\nIn recent decades, recurrent crises in world oil have resulted in serious economic crises, leading to the search for alternative fuels [1]. In 1930, Brazil presented the first National Congress on Industrial Applications of Alcohol that was aimed at establishing the infrastructure for the production and use of bioethanol. This led Brazil to start production in the early twentieth century, while other countries started the production of fuel from grains, using its potential for bioenergy production. The remainder of residues with high protein content is a source of nutrition in agriculture, as well as being a rich source of sustainability [2].
\nThe three major world powers producing bioethanol are Brazil (sugarcane), the USA (corn), and China (wheat and corn), where Brazil is the largest producer through a direct source of sugar, as production by grains requires an additional step with the liquefaction and hydrolysis of the starch. Estimates indicate that around 85% of all bioethanol worldwide is the responsibility of Brazilian and North American production, as well as inferior productions in all parts of the continents, as shown in Figure 1 [3].
\nGlobal distribution of production and percentage of production per continent: Americas: South America—Brazil 43% (sugarcane); North America: USA 44% (corn) and Canada 2%; Europe 3% (vegetables and wheat); Africa 1% and Oceania 1%.
In Brazil, the bioethanol had low volume of consumption compared to the use of conventional fuels, maintaining the Brazilian dependence on imported oil [4]. This made the national government launch the National Alcohol Program (ProÀlcool) in the 1970s, which established a new behavior for air quality and the development of technologies in the area of alternative sources of energy [5]. ProÀlcool represented the largest increase in bioethanol production, from 500 million liters at the beginning of the program to about 13 billion liters per year [6]. Since then, Brazil has been characterized as a potential producer of bioethanol, with a well-developed domestic consumption policy [6, 7].
\nCurrently, Brazil has an estimated bioethanol production with the 2016–2017 crops of 33.2 billion liters [8] and has kept that figure since 1986. All the production comes from sugarcane, representing a large-scale technology characterized by the development of new cane varieties, favorable climate, fertile soil, and advanced agricultural technologies [4, 6, 9]. The production of Brazilian bioethanol is derived from first-generation technologies, where a natural source of sugar from the sugarcane extraction, sucrose, is fermented by yeast with the primary product ethanol [10].
\nThe sugarcane plant used for the Brazilian bioethanol production is derived the crossing from 637 species of the genus Saccharum, family Poaceae, Andropogoneae tribe, and native of hot temperate climate and with morphology characterized by stem and straw [11, 12]. The stem is the material from which the sugarcane juice is derived and is later used for the production of sugar and bioethanol. The bagasse is composed of all post-grind materials and the trash, characterized by the dry, green leaves of the plants, which serve as products of fermentation in second-generation processes for the formation of bioethanol [13].
\nThe fermentation has been known since antiquity, being characterized as a biochemical and biological complex process, which has the objective of transforming sugar into ethanol (anhydrous and hydrated), carbonic gas, succinic acid, and volatile acids and esters [14].
\nThe Brazilian fermentation process is differentiated and unique due to the fact that it is fed-batch in most states, being these short fermentation cycles and cell treatments with sulfuric acid [10]. This process uses cane juice as raw material, with a final product of 9–12% (v/v) and an efficiency of 90–92% [15]. The ratio of bioethanol produced to the amount of raw material used varies according to the amount of sugar present in the must, which consists of a mixture of molasses (sugar manufacturing residue), water, and sugarcane juice. The process starts with an action of invertase exoenzyme, in the process of breaking the sugar (sucrose, a disaccharide) into glucose and fructose (structural monosaccharaides), which are absorbed by facultative aerobic microorganisms, which under anaerobic conditions form the pyruvic acid cycle, the enzymes pyruvate with the help of decarboxylase and alcohol dehydrogenase, producing the bioethanol and its subproducts at the end of the fermentation [9].
\nThe main key of the national fermentation process is that, at each end of the fermentation cycle, the yeasts are subjected to a centrifugation and sulfuric acid wash in order to minimize the risk of contamination [10]. At the end of this treatment, the cells are returned to the fermenters as a new inoculum for the subsequent cycle, this stage being repeated twice daily throughout the crop for 6–9 months, during the year, as shown in Figure 2 [16].
\nSimplified scheme of a fed-batch fermentation process with the recycling of yeast cells in Brazilian distillers by fermenting of sugarcane.
The fermentation with grains (the USA and China) is rich in carbohydrates so it is essential to the stage of liquefaction and hydrolysis of this raw material, where the molecules of starches are broken down into fermentable sugars, and thus fermentation can occur, as shown in Figure 3 [17]. One of the main characteristics of the grain fermentations, besides the additional stage of liquefaction and hydrolysis of the starch, is that the mills do not use recycled yeast cells, like the Brazilian mills, which is due to the fact that the whole concentration of residues and fermentable products is retained for distillation, decreasing the fermentation process when compared to the cell recycle process [18].
\nSimplified scheme of grains fermentation process with the liquefaction and hydrolysis in North American’s and Chinese’s distillers by fermenting of corns and wheat.
Significant changes are also observed when comparing the Brazilian and North American fermentation processes. In the fermentation of sugarcane, we have a lower concentration of solid residues, a concentration of larger yeast cells, and a much shorter time for bioethanol production, 6–12 h. The US process, that is derived for approximately 45–60 h, presents the advantages of a final concentration of bioethanol of 12–18%, against the 7–12% of the Brazilian process, and the raw material, that comes from corn plantations, lasts approximately one year, as opposed to the sugarcane harvest and its losses with rains that last around 200–240 day per year [19].
\nThe main microorganisms used for the fermentation process are yeasts, such as Saccharomyces sp., Schizosaccharomyces sp., Kluyveromyces sp., among others [13]. Currently, the most used yeast in the sugar and alcohol sector, for fermentation processes in the production of bioethanol is the specie Saccharomyces cerevisiae [20]. The methodologies used for the identification of yeasts based on morphology, biochemical characteristics, and sexual reproduction require the evaluation of 70–90 tests to obtain the identification of species. Macroscopic and microscopy features may be the first method of identification of S. cerevisiae yeasts, as presented in Figure 4 [21, 22].
\nMacroscopic and microscopy of yeasts Saccharomyces cerevisiae. Macroscopies (A), (B), and (C) of PE-2 isolates, grown in YEPD (yeast extract-peptone-dextrose growth medium) solid with creamy and yellowish-white culture characteristics. Microscopies: (D) clear field microscopy of PE-2 isolate; (E) PE-2 isolate with FITC (fluorescein isothiocyanate fluoroforo) and Propidium Iodide cell tags; (F) PE-2 isolate with Calcofluor cellular target; Microscopy presence of oval yeasts with budding presence, with size of approximately 4–8 μm. Microscopies were performed in IN Cell Analyzer, objective of 20× and diameter 70 μm.
The molecular techniques have been developed as alternatives to traditional techniques for the identification and characterization of yeasts, with the advantage of building an independent expression of the genes that allows quick and accurate identification of yeast species [23]. Due to the high mutation capacity of wild yeasts, molecular techniques for characterization and analysis of polymorphisms are being developed [24]. Genetic analyses of DNA, electrophoretic karyotyping, rRNA sequencing, rDNA restriction analysis, and polymerase chain reaction (PCR) and restriction fragment length polymorphism (RFLP) have been used as different tools to distinguish Saccharomyces sp. strains from strict sense group [25].
\nStudies of Melo Pereira [26] developed two new pairs of specific primers of the species, homologous to the HO gene of the species Saccharomyces bayanus, S. cerevisiae, and Saccharomyces pastorianus, offering a rapid method of PCR amplification, resulting in the correct identification of these species in less than 3 h. Guillamón [27] and Oliveira [28], by ribosomal DNA RFLP of ITS1, ITS2, and 5.8S identified different yeast species isolated from wine fermentation, and could also analyze the diversity of yeast species during spontaneous fermentation.
\nS. cerevisiae is characterized by being yeast with growth in media containing simple sugars and disaccharides, high genetic transformations, and qualities of high resistance to adverse conditions of the growth medium, offering a primordial role in the processes of fermentation [29]. Some strains of S. cerevisiae have the capacity to be highly productive, dominating the entire fermentation process during the harvest period, allowing efficient and stable fermentations, which result in lower costs and higher fermentation performance (high production capacity of ethanol), and high viability throughout the process [30].
\nStudies indicate that S. cerevisiae is adaptable to different environments, revealing to be a rich source of phenotypic profiles in the Saccharomyces sp. species evolution [31, 32]. It has recently been shown that the interaction between environmental factors and organism may influence the identification of different specific characteristics of S. cerevisiae [33, 34]. S. cerevisiae is widely used and cultivated in industrial fermentation, due to the high capacity of the yeast adaptations to the variable conditions of the environment, such as sugar and ethanol concentrations, pH, oxygen concentrations, resistance to contaminants, salt stress, protein stress, temperature changes, and osmotic pressure [35].
\nAccording to Gao et al. [36], using thermophilic strains is interesting in the processes that involve simultaneous saccharification and fermentation (SSF), as this process may reach from 45 to 50°C, resulting in a greater bioethanol production. This occurs because the yeast suffers less damage with the temperature increase and there is a lower chance of microbial contaminations. It is therefore desirable that the thermostable yeast fermentation occurs at the optimum temperature of the enzyme, maximizing the ethanol production process. Thus, the increase in thermotolerance in yeast results in cost production, increases yield in the ethanol production with simultaneous saccharification and fermentation (SSF) system and reduces the possibility of contamination.
\nHowever, it is important to know the fermentative yeasts for the control and monitoring of alcoholic fermentation, especially in search of selected characteristics of dominance and resistance to bioethanol yeast production. The objective of this chapter is to distinguish the main phenotypic characteristics of S. cerevisiae yeasts in the alcoholic fermentation, for a possible selection of new strains with differentiated phenotypic characteristics, resistant and ideal for the production of bioethanol.
\nCell flocculation of yeast strains such as S. cerevisiae is called cell aggregation and sedimentation in liquid media [37]. Cells have the characteristics of agglomeration at the end of each fermentation process, which makes it an interesting and divergent phenomenon in the industry, as shown in Figure 5 [38]. Studies point out some divergences in flocculation, which can be a phenomenon of cooperative protection mechanism found in cells during adverse factors in the fermentation cycle [39]. It also facilitates the separation of yeasts at the end of the fermentation by sedimentation, thus helping the collection, centrifugation, and cellular treatment, contributing to the new inoculum stage, for a next cycle in the fermenter [40].
\nCharacterization of aggregative power of Saccharomyces cerevisiae yeast cells, grown in YEPD liquid medium and stained with lactophenol. (A) Cells in normal condition without aggregative power. (B) Cells in condition of cellular aggregation.
The principle of cell adhesion is initiated through the recognition of mannose chains, located on cell surfaces, by lectin-like proteins, and agitation is necessary for the beginning of flocculation [41]. Two hypotheses for flocculation are well established: (I) sensitively to proteinases; (II) inhibited by saccharides, suggesting the existence of a protein that recognizes these sugars [42].
\nThese two hypotheses classify two flocculation groups in yeasts that are distinguished by the inhibition of sugar: the first group, called New Flo phenotype, characterized by the inhibition of mannose, glucose, maltose, and sucrose with the exception of galactose; and the second group, the Flo1, is inhibited by mannose, but not by glucose, maltose, sucrose, and galactose, and its action is normally bound to a gene [43].
\nIt is believed that these two distinct phenotypes are caused by two different proteins of the lectin type. Furthermore, the physical-chemical interaction in the cells surface may be involved in the aggregation process, where there is a correlation between flocculation and electrophoretic mobility of yeast cells on certain stress conditions. Other studies reported the correlation of hydrophobicity in the process of flocculation [44].
\nAnother hypothesis for flocculation is the action of a dominant gene family (FLO1, FLO5, FLO8, FLO9, FLO10, and FLO11), where they encode a yeast cell wall protein that acts directly on the cell aggregation [45]. The proteins encoded by these FLO genes share a cellular/modular organization in three domains: an amino-terminal responsible for carbohydrate binding, a central domain, and a carboxyl-terminal domain containing a glycosylphosphatidylinositol anchor sequence [46]. However, the central domain contains tandem repeat regions of DNA sequence that can drive recombination reactions within and between FLO genes, resulting in new generations of FLO alleles, thus conferring yeast cells a wide diversity in the flocculation phenomenon [47].
\nThe environmental adversities occurring in a fermentation cycle, such as the decrease of nutrients by sugar consumption, temperature changes, pH changes, risk of contamination, phenolic compounds, and the concentration of ethanol by its own production occurs in different forms and some of them were completely studied [48]. An understanding of the cellular mechanisms of protection to the multiphysical and chemical stresses that the yeast undergoes during fermentation cycle is fundamental for the selection of ideal yeast [49].
\nTemperature elevations result in reduced fermentation efficiency in S. cerevisiae, due to the high fluidity in the membranes, caused by the altered composition of fatty acids in the adverse response [50]. As one of the stress factors known in the fermentative cycle in yeast, temperature change restricts ethanol production and induces the accumulation of proteins bound to tolerance stress [51].
\nIn the first-generation fermentation cycle, yeasts require a temperature of 30°C, whereas, in the production of second-generation bioethanol, where cellulose enzymes start the process by saccharification, yeasts require a higher temperature of 45–50°C [52]. The efficacy of the fermentation is decreased at high temperatures, because it causes damage to the yeast cell, such as the rupture of the protein structure or the loss of function, thus preventing cell proliferation, decreasing viability during the process, and leading to cell death [53]. This temperature control in the fermentative cycles is a problem for the plants in tropical countries, where the ambient temperature is already naturally high and cooling systems are necessary for the total control of this temperature [54].
\nIn the bioethanol production, the process temperature must be stabilized at around 30°C (the cell growth temperature), which is reaching 40°C [55, 56]. Thus, thermotolerant yeast strains may be a promising approach to a profitable fermentation process, as is the case of simultaneous saccharification and fermentation that requires high temperatures to increase ethanol yield [36].
\nOsmotolerance can be an important factor in the production of ethanol for its adaptation strategy employed in all cell types by accumulating compatible solutes (sulfite), resulting in a decrease in the potential of intracellular water [57]. As sulphite and sulfite-generating compounds have long been used as antimicrobial agents in alcoholic fermentation, tolerance to sulfite in yeast is another desired characteristic for the production of bioethanol from sugarcane juice [58].
\nThe high levels of ethanol in the fermentation medium are considered as negative parameters in the process conditions, because at the same time that the production is essential, the accumulation of ethanol by this production generates an acidification of the medium, leading to irreversible damages in the yeast membrane, thereby decreasing cell viability [59].
\nThe true physiological and ecological relevance of ethanol tolerance in S. cerevisiae is its ability to generate mechanisms that protect the cell from chemical and physical damage at high levels of ethanol [60]; this is usually observed in a typical fermentation environment, where there is a large amount of sugars, leading later to ethanol production [30]. This stage generally occurs by stationary phase cells and its tolerance to the ethanol produced is only controlled by the integrity of the yeast membrane in contact with the ethanol accumulation, which is composed of chitin, glucans, glycoproteins, fatty acids, and ergosterol [61].
\nHowever, S. cerevisiae is resistant to ethanolic stress for its capacity of modifying the conformation of its membrane in the increase of fatty acids and ergosterol when coming in contact with the adverse environment, thus neutralizing the damages caused, mainly in relation to its viability [62, 63]. The accumulation of ethanol can also affect the structural compliance of the cellular proteins causing the inefficiency of its actions, such as the decrease of the activity of glycolytic enzymes: pyruvate kinase and hexokinase, besides altering the absorption of glucose, maltose, and amino acid. In some cases, there may occur cellular extravasation of essential cellular components [64].
\nIn industrial fermentations, a high capacity of production is observed by the accumulation of ethanol in the medium, indicating a positive assimilation of residual sugar, which is measured by the visualization of cellular proliferation in the presence of the gradual levels of ethanol produced during the fermentative process [65]. Tolerance and ethanol characteristics of the main industrial strains of S. cerevisiae studied are described in Table 1 [30, 66–69].
\nStrains | \nGroup | \nOrigin | \nFeedstocks | \nTemperature tolerance (°C) | \nEthanol tolerance (%) | \n
---|---|---|---|---|---|
ZTW1 | \nIndustrial (fuel ethanol) | \nChina | \nGrains | \n55 | \n18 | \n
YJ5329 | \nIndustrial (fuel ethanol) | \nChina | \nGrains | \n55 | \n18 | \n
PE-2 | \nIndustrial (fuel ethanol) | \nBrazil | \nSugarcane | \n40 | \n15 | \n
CAT-1 | \nIndustrial (fuel ethanol) | \nBrazil | \nSugarcane | \n40 | \n15 | \n
AT-3 | \nIndustrial (fuel ethanol) | \nUSA | \nGrains | \n40 | \n14 | \n
ErOh red | \nIndustrial (fuel ethanol) | \nUSA | \nGrains | \n40 | \n15 | \n
Characteristics of temperature tolerance and ethanol for major industrial strains worldwide used for the production of bioethanol.
Although this assay is routinely used in industries as large-scale screenings, its actual importance in ethanol resistance in yeast is not elucidated, due to divergent of actions that this process can cause, for example, the negative side acting in the decrease of the cell viability, and positive the increase in resistance to contaminating microorganisms in the fermentation process [70].
\nThe metabolic pathways correlated to the expression of genes responsive to high levels of heat stress and ethanol stress include heat shock proteins (HSPs) and also metabolic enzymes such as trehalose, which is directly involved in tolerance in S. cerevisiae [71]. HSPs play a role in folding and refolding, transport, and degradation of intracellular proteins, triggered by stress in fermentation process and located in the cytoplasm, nucleus, and mitochondria, acting immediately in response to an accumulation of denatured proteins, activating the transcription factors of thermal shock (HSF), and leading to a positive regulation of thermotolerance gene expression [72].
\nThe interactions of multiple genes at loci for cellular functions under heat and ethanol stresses are essential [73]. HSPs are known as chaperones ensuring the functional and structural conformation of the yeast, on the action of genes such as SSA1, SSA2, SSA3, and SSA4 which are expressed together with the HSP genes HSP12, HSP26, HSP30, HSP31, and HSP150 which were also found active at high stress levels [74] and interactions between chaperones of different types are widely encountered [75].
\nHowever, the inference of several chaperones shows an effective activity in neutralizing the stress, with the activation of the functional chaperones specific to more complex structures in the yeast cell walls [76], which have as a main function to repair of these denatured proteins to maintain cell viability [77].
\nIn addition to serving as chaperones, HSPs have numerous other functions, for example, Hsp30p is characterized as a hydrophobic plasma membrane protein that acts on the regulation of H+-ATPase, Hsp31p, and Hsp32p functions as hydrolases and peptidase, and Hsp150 is characterized as a protein in supporting the cell wall stability and remodeling [78]. HSPs and chaperone-mediated genomic regulation are also linked to glucose metabolism, which are indispensable tools for stress tolerance in yeast metabolism, especially with storage of carbohydrates, such as trehalose [79].
\nTrehalose is a compound that acts to prevent the influx of excess salts resulting in irreversible dehydration of cells; therefore, yeasts are capable of accumulating trehalose up to 15% in a stress environment [80]. The trehalose acts by reducing the permeability of the membrane thereby rendering it hydrophobic, due to some regulatory genes such as TPS1, TPS2 and, TSL1, as well as acting in the remodeling of proteins under stress conditions [81]. Cells incapable of accumulating trehalose presented depreciated growth, leading to a significant decrease in cell viability during fermentation stresses [82].
\nSugarcane juice is one of the main means used in the production of bioethanol, which is derived from the break of fermentable sugars such as sucrose, glucose, and fructose in contact with fermenting microorganisms such as yeast S. cerevisiae [83]. Yeast consumes the sugars in the medium in a complex and highly regulated manner, the principle of fermentation, where the sucrose is consumed first, followed by glucose and fructose, and finally maltose, this assimilation of sugars can occur simultaneously between the breaks of sugars, which is the standard process for sequential uptake of the glucose repression pathways or the catabolite repression pathway [84].
\nGlucose and sucrose may trigger beneficial effects on cells, including stimulation of cell proliferation, mobilization of storage compounds such as glycogen and trehalose, as well as decreased resistance to cell stress [85]. In contrast, negative impacts due to lack of glucose in the process can lead to several problems such as decreased or blocked fermentations, instability of cellular viability and low ethanol production [86], where the break of sugars, sucrose into simple sugars (glucose) occurs by an intracellular enzyme known as invertase, located in wall the yeast industries [87].
\nMicroorganisms that possess the ability to assimilate the highest amount of sugars are indicated for the production of bioethanol, examples are shown in Table 2 characterizing the main strains of S. cerevisiae worldwide used in industries for the production of bioethanol [30, 66–69].
\nStrains | \nGroup | \nOrigin | \nFeedstocks | \nAssimilation sugar (%) | \nProduction bioethanol (%) | \n
---|---|---|---|---|---|
ZTW1 | \nIndustrial (fuel ethanol) | \nChina | \nGrains | \n65 | \n28 | \n
YJ5329 | \nIndustrial (fuel ethanol) | \nChina | \nGrains | \n60 | \n33 | \n
PE-2 | \nIndustrial (fuel ethanol) | \nBrazil | \nSugarcane | \n51 | \n22 | \n
CAT-1 | \nIndustrial (fuel ethanol) | \nBrazil | \nSugarcane | \n52 | \n26 | \n
AT-3 | \nIndustrial (fuel ethanol) | \nUSA | \nGrains | \n42 | \n18 | \n
ErOh red | \nIndustrial (fuel ethanol) | \nUSA | \nGrains | \n75 | \n30 | \n
Characteristics of the assimilation of residual sugars and ethanol production for large industrial strains of Saccharomyces cerevisiae yeasts used in the production of bioethanol.
The assimilation of sugars in the fermentation process is not exclusively the fermentation of sugarcane. Currently, new technologies are available to produce ethanol from vegetables such as potatoes, cassava, beets, cereals such as corn, and there are also studies showing the production of green bioethanol in algae fermentations [88].
\nThis type of fermentation is due to the breakdown of starch, carried out by the action of the enzyme glucoamylase, acting directly on the conversion of starch to glucose, by breaking the successive bonds of the nonreducing end of the glucose finally producing straight chains [89]. The process of producing ethanol from starch involves two main steps: enzymatic hydrolysis as the main step and habitual fermentation as the second step [90].
\nAll adverse parameters studied for the first-generation fermentation process have been highly researched to reach an ideal model of production of second-generation bioethanol, which is characterized by being profitable and environmentally sustainable [91].
\nSecond-generation bioethanol production starts from the lysis of the raw material (sugarcane bagasse, vinasse, and residues from the milling of grains). The main step is characterized by a pretreatment where the breakdown of the cellulose-hemicellulose-lignin complex allows the production of fermentable sugar levels for a subsequent fermentation, demonstrated in Figure 6 [92].
\nSimplified scheme of second-generation fermentation process with steps of biomass and hydrolysis for bioethanol production worldwide.
The hydrothermal and lime pretreatments are the most used, known for making the method more effective in preparing the biomass bioconversion step [93], a strong advantage for the sugarcane bagasse. They can be carried out under conditions of low temperature and pressure, resulting in lower sugar degradation, whereas in the saccharification the pretreatment is observed with high temperatures and difficult breaks of carbohydrate chains, resulting in a lower amount of sugars [94].
\nThe fermentation of lignocellulose hydrolysates for bioethanol production presents two main problems: first, the fermentation of xylose that requires a low and controlled oxygenation; second, the removal of microbial inhibitors, which can contaminate the process [95]. Furthermore, these yeasts present a certain tolerance limited to ethanol [96].
\nGenetic manipulation in the metabolism of xylose in yeast fermentation has advanced and pioneering studies on glucose transporters that mediate xylose uptake, allyl-xylitol-reductase genes, xylitol dehydrogenase and xylulokinase have been expressed, which allows a better assimilation and fermentation of xylose [97].
\nThe main concern in this step is that the balanced supply of NADP (enzyme nicotinamide adenine dinucleotide phosphate) and NADPH (enzyme nicotinamide adenine dinucleotide phosphate oxidase) has to be constant to avoid the production of xylitol. The path is the reduction of NADPH production by blocking the oxidative pentose phosphate cycle in xylose assimilation [98]. Cellulose hydrolysates present different inhibitors from lignin derivatives and sugar degradation, resulting in high amounts of acetic acid, intrinsically necessary for the deconstruction of biomass [99].
\nAll adversities of typical fermentation first generation associated with pH, temperature, elevation of ethanol concentrations, and temperature among other stress factors present in large-scale fermentations are seen together in the adversity challenges of second-generation fermentation [100].
\nFirst-generation fermentation over the years has mainly been used for large-scale industrial models. Although it is a well-established process, it is not definitively elucidated. Changes can be seen with each new process initiated presented for fundamental parameters and the behavior of the yeasts used.
\nS. cerevisiae has the characteristic of being adaptable to any environment, which leads to numerous behavioral responses during fermentation. As for each new cycle, changes are inevitable and checking all parameters of fermentation are of extreme importance for the success of the fermentation.
\nFlocculation is a divergent parameter, although it can have many advantages as a phenomenon of cellular protection to several stressors and contaminations in a process, it also presents disadvantages such as low yield in fermentation of fermenting tanks by their decanting. The question whether this phenomenon is beneficial or detrimental on flocculation is still uncertain; however, it is well-known and elucidated in its morphology or molecular action in yeast cells, and it contributes to the improvement of bioethanol production in the world industry.
\nFor an alcoholic fermentation to be efficient, it is necessary and indispensable to know what happens throughout the process, the main steps and degrees that microorganisms go through for hours and days in order to remain viable and productive. For this reason, the study and knowledge of the two main parameters stress of fermentation (heat and ethanol) is of paramount importance for any beginning of the process, whether in small scale, as in laboratories, or large scale, as in industrial productions. The behavioral responses of the fermentation are measured through these parameters that are observed at all times, always aiming the improvement for the process.
\nThe main step for a virtuous bioethanol production is the ability of the microorganism to breakdown the sugars and thus assimilate them to ferment. This detailed step has to be well studied so that there is no damage throughout the process, especially at industrial scales, so that both, a sufficient amount of microorganism concomitantly and adequate amount of sugars are essential to the start of the production.
\nLooking at the current scenario, the first-generation processes were modernized and studies and improvements resulted in second-generation fermentation, which aims to take advantage of all remaining residues and reaches to more sustainable processes. These processes are taking strides and improvements are being seen at all times to reach the ideal process.
\nIn view of this profile, the search for yeasts with more robust characteristics in industrial lines is essential, and different strategies involving adaptation and functionality are highlighted by genetic engineering research. Advances in the area of a process and ideal yeast are positive, but the journey is still far from reaching perfection. The secrets and mysteries of fermentation are innumerable, but research is constantly revolutionizing and little by little these are being unraveled and the beginning of everything is the understanding of all the steps and all its parameters.
\nThis study was financially supported by PADC-FCFAR UNESP and scientific agencies CAPES and CNPq.
\nChlorophyll and carotenoid are important pigments that have been used as intrinsic optical molecular probes to observe plant performance during different phases of development. Chlorophyll and carotenoid are biosynthesized in chloroplast and their metabolism is closely related with the chloroplast development. Chlorophyll biosynthesis begins with the formation of 5-aminolevulinic acid (ALA) from glutamate (Glu) via Glu-tRNA synthetase, Glu-tRNA reductase (GluTR) and Glu-1-semialdehyde aminotransferase (GSA-AT) [1]. Eight molecules of ALA are condensed, eventually forming the symmetric metal-free porphyrin, protoporphyrin IX (Proto IX), which is a common precursor of haem and chlorophyll. The biosynthesis of chlorophyll continues by insertion of Mg2+ into Proto IX and followed by several steps in the chlorophyll cycle to create protochlorophyllide.
\nFurther, reaction is one of the most interesting steps because this is the first step in chlorophyll biosynthesis that requires light: the NADPH:protochlorophyllide oxidoreductase converts protochlorophyllide into chlorophyllide. This reaction is then continued to produce chlorophyll (chl) a and b. So, when dark-grown etiolated seedlings are exposed to light, protochlorophyllide is immediately converted to chlorophyllide and then further to synthesis of chl. Once chl a and b are formed and properly incorporated into the thylakoid membranes and associated photosystems, chloroplast is fully functional to do photosynthesis [2].
\nPlant carotenoids are synthesized and accumulated exclusively in plastids, most importantly, chloroplast and chromoplast [3]. There are two types of plant carotenoid: carotene, which is cyclized and uncyclized hydrocarbons, and xanthophylls, which are oxygenated derivatives of carotenes. Carotenoid synthesis is initiated by the formation of C40 compound phytoene by the head-to-head condensation of two molecules of geranylgeranyl diphosphate (GGDP) by phytoene synthase and then to a series of 4 sequential desaturation reactions, by two separate enzymes to produce lycopene, which has 11 conjugated double bonds [4]. Lycopene is then cyclized to α-carotene or β-carotene, which is then further hydroxylated to produce colorful xanthophylls such as lutein, β-cryptoxanthin, zeaxanthin, antheraxanthin, violaxanthin and neoxanthin. The biosynthesis and accumulation of carotenoids in the dark-grown etiolated seedling are essential for the assembly of membrane structure and benefits the development of chloroplast when seedlings emerge into the light [5]. Understanding the relationship between structure and photophysical properties of these pigments can provide insights into a better study of how photosynthesis works at the molecular level in chloroplast.
\nThe photophysical properties and functions of chlorophyll and carotenoid reside in their chemical structure. Chlorophylls are defined as cyclic tetrapyrroles carrying a characteristic isocyclic five-membered ring that are functional in light-harvesting or in charge separation in photosynthesis [6]. The chemical structure with IUPAC numbering scheme of chl a is shown in Figure 1. It is a squarish planar molecule, about 10 Å on a side. An Mg atom in the center of the planar portion is coordinated to four nitrogen atoms. The five rings in chlorophylls are lettered A through E, and the substituent positions on the macrocycle are numbered clockwise, beginning in ring A. Chlorophyll has two molecular axes: y-axis is defined as passing through the N atoms of rings A and C and x-axis passing through the N atoms of rings B and D. The delocalized π electron system extends over most of the molecule, except for ring D, in which the C-17—C-18 double bond is reduced to a single bond. The tail is formed by condensation of four isoprene units and is then esterified to ring D. It is often called phytol tail, after the polyisoprenoid alcohol precursor that is attached during biosynthesis. Because of the reduced ring D, plant chlorophylls such as chl a and b are classified as chlorins rather than porphyrins. These types of pigments have (in organic solvents) absorption bands around the blue and red spectral regions (Figure 2a), which are called B (or Soret) and Q bands, respectively, and arise from π→π* transition of the four frontier orbitals [7, 8]. One band each pair is polarized along the x-axis (Bx, Qx) and other along y-axis (By, Qy). The strong absorption band at the maximum absorption wavelength (λmax) 660 nm is called Qy transition band, which corresponds to the electronic transition polarized along the y-axis. The Qx-band of chl a shows a weak band near 550 nm, while the two overlapping Soret (B) bands show at about 430 nm. The chemical structure of Chl b is identical to chl a except at the C-7 position, where a formyl group replaces the methyl group. This structural change results in a shift of the Qy maximum absorption to shorter wavelength. The fluorescence spectrum of chlorophylls peaks at slightly longer wavelengths than the absorption maximum. The fluorescence emission (Figure 2b) is polarized along the y molecular axis, as it is emitted from the Qy transition. Shift of the emission to the longer wavelength side of the main transition is known as Stokes shift. In light reaction, chlorophyll plays as key pigment in the collection of light energy in the light-harvesting complexes and to carry out reversible photochemical redox reaction (Krasnovsky reaction) in the reaction centers.
\nChemical structure of Chl a (a), Chl b (b), lycopene (c), β-carotene (d), zeaxanthin (e) and lutein (f) with IUPAC numbering system.
(a) UV–Vis absorption of Chl a (black) and Chl b (red) in MeOH, (b) fluorescence emission spectra of Chl a (red) and Chl b (black) in MeOH, (c) β-carotene (red), zeaxanthin (black) and lutein (blue) in EtOH, (d) lutein in several organic solvents; MeOH (black), acetone (pink), diethyl ether (purple), hexane (light blue), EtOH (blue).
Structure of carotenoid is characterized by a linear chain of conjugated π-electron double bonds (Figure 1). In oxygenic organisms, carotenoid usually contains ring structures at each end, and most carotenoids contain oxygen atoms, usually as part of hydroxyl or epoxide groups. The primary molecular factor that gives rise to their strong absorption bands in the visible spectral region is the number of π-electron conjugated double bonds, N. The position of the absorption maxima is affected by the length of the chromophore, the position of the end double bond in the chain or ring and the taking out of conjugation of one double bond in the ring or eliminating it through epoxidation. Progressive movement to longer wavelengths (bathochromic shift) is illustrated by the absorption spectra of the acyclic carotenoid of increasing chromophore length. Carotenoids show different optical characteristics in various solvents, depending on the polarizability of the solvent [9, 10]; however, generally they have a typical three-peaked absorption spectrum with well-defined maxima and minima (fine structure) (Figure 2a). A ring closure as in β-carotene produces a less-defined fine structure. The introduction of a carbonyl group in conjugation with the polyene system produces a bathochromic shift and the loss of fine structure [4]. The influence of other substituents such as OH is negligible, for example, β-carotene, cryptoxanthin and zeaxanthin all have very identical absorption spectrums. Owing to the double bonds in the molecule, all carotenoids exhibit cis-trans isomerization (stereomutation). A cis double bond implies a configuration with the highest-priority group on the same side, whereas in the trans configuration they are on opposite sides. The absorption spectrum of a cis isomer presents a subsidiary peak in the near-ultraviolet, the cis peak; generally, it is located 143 nm from the longest wavelength maximum. For example, cis peak will appear at 330 nm if the longest wavelength maximum is 473 nm. In photosynthetic systems, carotenoid has essential functions. First, carotenoid is an accessory pigment in the collection of light energy in the spectral region which chl does not absorb and in transferring energy to a chl pigment [11, 12]. Second, carotenoid functions in a process called photoprotection by quenching triplet state of chl before it reacts with oxygen to form singlet oxygen species (ROS) [13, 14]. Third, carotenoid regulates energy transfer in the light-harvesting antenna through a process called xanthophyll cycle, to avoid over-excitation of the photosynthetic system by safely dissipating excess energy [15, 16].
\nIn the chloroplast interior, there are four main constituents in plant thylakoids, that is, photosystem II (PSII), cytochrome b6f, photosystem I (PSI) and the ATP synthesis. Chlorophylls and carotenoids are embedded in PS II and PSI, large pigment-protein clusters, the structures of which are perfectly adopted to ensure that almost every absorbed photon can be utilized to drive photochemistry. Both PSII and PSI consist of two moieties, that is, core complex or the reaction center that is responsible for charge separation and light-harvesting antenna complexes that surround the core complex and have functions to increase the capture of light energy and energy transfer to the reaction center in the core complex.
\nOne can detect chlorophyll and carotenoid bound in PSII and PSI in chloroplast by measuring their absorption and fluorescence spectra. Figure 3a (solid red line) shows the absorption spectrum of diluted chloroplast that is indicated by red shift of Chl a, Chl b and carotenoid’s bands because these molecules are bound as pigment-protein complexes in chloroplast. The Soret band of chl a in the complexes was detected at 438 nm while in the MeOH it was found at 432 nm (Figure 2a black line). The fluorescence emission spectra (Figure 3b) indicate a strong emission band of PSII complexes with maximum wavelength at (λmax) about 682 nm and weak emission band of PSI complexes with λmax at about 730 nm. It is shown here that Chl a acts as the main contributor to the excitation band at 434 nm and it shows that excitation at 434 nm (Soret band) produces stronger emission intensity, while the excitation at 475 and 512 nm, correspond to Chl b Soret band and carotenoid, respectively, produces weaker emission intensity. If we monitor the emission at 682 nm and measure the excitation spectrum, it shows that the PSII emission at 682 nm is the result of contribution from Chl a, Chl b and carotenoids (Figure 3a solid black lines) with bands at λmax about 414, 434, 475 nm, respectively.
\n(a) Overlaid of UV–Vis absorption (red) and fluorescence excitation (black) (λem = 682 nm) spectra of chloroplast and (b) emission spectra of chloroplast with excitation at 434 (black), 475 (red) and 512 (blue) nm. Measurements were conducted at ambient temperature. The isolation of chloroplast was carried out as follows: 20 g of suji leaves (Pleomele angustifolia) were washed with running water and cut. The leaves were then homogenized in 200 mL ice-cold isolation buffer (300 mM sorbitol, 50 mM HEPES-KOH pH 7.5, 2 mM EDTA, 80% acetone, 0.1% BSA) for 10 min in a cold environment, followed by filtration using cloth. Centrifugation was conducted in 2 steps, to discard cell debris at 200 g, 4°C, 20 min and to harvest chloroplast pellet at 3000 g, 4°C, 20 min. Final chloroplast pellet was collected and subjected to spectrum UV–VIS (Shimadzu UV-1700)and fluorescence measurement (Jasco FP-8500).
The current high-resolution structural models of antenna complexes have been obtained only for LHCII (2.72 Å) and recently for CP29 (2.8 Å) from PSII of spinach [17, 18]. Here we focus more on the LHCII structure. LHCII shows trimeric structure. Each monomeric contains three transmembrane α-helices, a, b and c (Figure 4a). One monomeric subunit contains eight chlorophyll (Chl) a pigments, six Chl b, two luteins (Lut), neoxanthin and one additional xanthophyll [17, 19]. The 14 chlorophylls are non-covalently attached in the protein cavity. Four carotenoid binding sites per monomer have also been characterized, but in this case the type of carotenoid bound can vary. Typically, two lutein molecules are in groves on both sides of helices a and b and have been likened to a cross-brace. A third carotenoid, 9-cis neoxanthin, is located in the Chl b-rich region near helix c. The fourth carotenoid is located at monomer-monomer interfaces in the trimer. It has been suggested that this site accommodates carotenoids that can participate in the xanthophyll cycle. It depends on the external stress level of the plant; the fourth carotenoid is either violaxanthin (no or low stress) or zeaxanthin (high stress) [20]. In this structure, the carotenoids are in van der Waals contact with the chlorophylls [9]. This is essential as carotenoids in LHCII act as accessory light-harvesting pigments and photoprotectors. The accessory light-harvesting function represents singlet-singlet energy transfer from the carotenoid to the chlorophylls. Since the singlet excited state lifetime of the carotenoid is quite short, approximately 200 fs, the carotenoid must be in close distance to a chlorophyll molecule if the energy transfer is to be efficient. Photoprotection function represents the quenching of triplet excited state of chlorophylls and so preventing the formation of singlet oxygen. This triplet-triplet exchange reaction also requires the carotenoid to be in close contact with the chlorophylls. Regarding CP29, it binds 3 carotenoids and 13 chlorophyll molecules [18]. The position of some chlorophyll binding sites in CP29 differs from LHCII.
\n(a) A view looking down on the top of trimeric complex of LHCII structure from spinach. Each monomer is colored magenta, yellow and pale green. The three-transmembrane helices (a, b and c) present in a monomer are labeled and are easily visible. Chl a molecules are in red, Chl b colored green and carotenoids colored orange. (b) Side-view of LHCII structure shows chlorophyll and carotenoid molecules are packed densely and close to each other (within van der Waals contact), enabling the crucial photo-protective role of these molecules to function by quenching triplet chlorophyll excited states. (c) Structure of PSII from Thermosynechococcus elongatus [28], a side-view representation of the overall dimer perpendicular normal with the pseudo-twofold symmetry axis. (e) PSII core reaction center is shown; component co-factors of the electron transport chain viewed along the membrane plane. The two branches are related by the pseudo-twofold symmetry axis. The respective pairs of pigments on the branches are labeled to indicate whether the Mg2_ is coordinated by D1, D2. (d) Structure of PSI from Synechococcus elongatus [29]; overview of the complete trimer looking along the membrane normal from the stromal side with each polypeptide of the trimer colored differently and chlorophyll molecules given in green. The two main proteins that comprise a monomer are PsaA (yellow) and PsaB (magenta). The electron transport chains are in the center of each monomer. (f) PSI core reaction center component co-factors of the electron transport chain are viewed along the membrane plane. The two branches are related by the pseudo-twofold symmetry axis. The representative pair of chlorophyll molecules on the branches are labeled A or B indicating whether the Mg2+ is coordinated by PsaA or PsaB. The iron–sulfur center of Fx involves residues from both PsaA and PsaB, while FA and FB are located in an extrinsic subunit called PsaC.
The current high-resolution crystal structure of PS II and PSI core complexes is limited to that from cyanobacteria and from pea, respectively [21, 22]. The core of PSII is a multi-subunit complex. Most of the chromophores involve light harvesting as well as electron transfer reaction and are bound to four main subunits, that is, D1, D2, CP43 and CP47. When the core of PSII and PSI reaction center structures is compared, the arrangement of the pigments and other electron transfer co-factors is also very similar (Figure 4c and d). Here, first we look at the PSII core reaction center. The core of reaction center of PSII is made from two major polypeptides called D1 and D2; each contains five membrane-spanning α-helices. These two helices clasp each other like two cupped hands holding on to each other. The redox cofactors are arranged into two arms that lie on either side of the point where the two groups of helices interact. This arrangement of the helices and the cofactors introduces a pseudo two-fold symmetry axes that runs through the center of reaction center normal to the plane of the membrane. In Figure 4e, it is seen that the electron transport pathway in PSII begins with a pair of chlorophyll molecules called P680 (PD1 and PD2). Then each arm contains, in order, one monomeric chlorophyll molecule, one pheophytin (a chlorophyll derivative) and one plastoquinone molecule. Here, only the D1 arm is active in electron transport. Upon excitation P680 becomes oxidized and one electron is injected out and passes down the active branch to the quinone QA. P680 is re-reduced by electron transfer from a special tyrosine residue called Z (Tyrz). A second turnover of P680 delivers a second electron to the plastoquinone and the secondary quinone QB is now reduced to QBH2. The hole on Tyrz is filled by electron transfer from the manganese cluster, the oxygen evolving complex. Every four turnovers of P680 stores four positive charges in the manganese cluster that are then used to oxidize water and evolve oxygen. While in CP43 and CP47, there are a total of 49 Chl a molecules that are bound and that function as internal antenna and allow excitation energy transfer from the peripheral antenna system to the reaction center.
\nUnlike PSII, in PS I, the same single polypeptides contain both antenna complexes (Lhca) and the reaction center core. The 3.3 Å resolution crystal structure of PSI from pea showed that plant PSI binds at least 173 Chl a and b molecules [22]. At this resolution of the crystal structure, it is not possible to identify the Chl species, but biochemical analysis of purified PSI indicated that it has a Chl a/b ratio in a range of 8.2–9.7 [23, 24]. A large number of Chl a and b molecules are bound to the Lhca protein, only about 100 Chl a are bound in the core complex, and the rest of Chl a and b are between these moieties. The latter represent the so-called “linker” chlorophylls which are located between Lhca monomers and “gap” chlorophylls (between Lhca and PSI core). The linker chlorophyll molecules probably play an important role in excitation energy transfer between Lhca antennas and from Lhca to the PSI core [20, 25, 26]. Based on biochemical analysis, PSI was reported to bind approximately 33/34 carotenoids, that is, about 12 carotenoid molecules are bound to Lhca, at the interface between Lhca and the core complex, and about 22 β-carotene are bound to the core [20, 23, 26]. Based on these biochemical analysis, it can be estimated that PSI-LHCII supercomplex contains about 215 chlorophyll and 45/46 carotenoid molecules.
\nThe core complex of PSI is composed of smaller number of subunits (15 subunit) than PSII. The large PsaA and PsaB subunit, which contain 11 trans-membrane helices each, forms a hetero-dimer that binds ~80 Chl a and ~20 β-carotene as cofactors for light harvesting as well as 6 Chl a, 2 phylloquinones and a 4Fe-4S cluster as cofactors for electron transfer reaction, with the exception of terminal electron acceptors (Fe-S clusters FA and FB) which are bound to the PsaC subunit [25]. At closer look (Figure 4f), the redox co-factors in the core reaction center are arranged into two arms that are located on either side of the region where two groups of helices interact with each other. Two chlorophylls form P700 and then each arm contains two monomeric chlorophyll molecules (the second one being in the equivalent position to the pheophytin present in photosystem II) followed by one quinone molecule. When P700 is oxidized, both arms of the electron transport pathway are able to work as it was reported that the electron can pass either down the B-branch or the A-branch [27].
\nChlorophyll and carotenoid can be isolated as free pigments, detached from the pigment-protein complexes, by organic solvent extraction. Important aspects such as the choice of organic solvents, light exposure and working temperature should be considered while isolating pigments. Based on the structure, chlorophyll is characterized with polar macrocycle ring with non-polar hydrocarbon tail. The structural difference between Chl b and Chl a is by having an aldehyde group in place of the methyl group at the macrocycle side group. This change is effecting the polarity of Chl b to be more polar in comparison to Chl a. In the case of carotenoid, structural difference can be seen from the number of conjugated double bonds and the presence of oxygen atoms. Considering these characteristics, mixtures of miscible polar and semi/non-polar solvents are used commonly to extract plant pigments. The mixture of solvent has double functions, that is, penetrating tissues/matrixes and extracting pigments from their lipophilic surrounding. During extraction, exposure of light should be avoided to reduce photodamage of the pigments. Temperature is also important. It is recommended to conduct extraction at lower temperatures, for example, on ice or using liquid nitrogen, to minimize activity of enzyme (e.g. chlorophyllase) that will catalyze breakdown. Antioxidant agent can be also added during extraction to avoid any unwanted oxidation.
\nAfter successful isolation, liquid chromatography has been widely used as an effective technique to separate individual type of pigments and for further purification. In this technique, the pigment separation is based on the polarity which depends on the interaction of pigment with the stationary and mobile phases. Elution method either normal phase or reversed phase is chosen according to the type of pigment to be separated. In addition, the choice of liquid chromatographic methods, namely thin layer chromatography (TLC), column chromatography (CC) and high-pressure liquid chromatography (HPLC), is referred to the speed, resolution and quantity of sample [30]. Currently, ultra-fast liquid chromatography (UFLC), a recent development of HPLC, has been used as a standard for liquid chromatography to achieve high-resolution data with low time consumption [31]. Purification with non-chromatographic method has also been developed, that is, purification method using dioxane has been effective to separate chlorophyll from most of the carotenoids and some lipids [32].
\nVarious types of column absorbents used for chromatographic separation of plant pigments have been well reviewed [30]. Here, we used a silica C30 column attached to UFLC analytic to achieve well separation of carotenoids from Pleomele angustifolia leaf using elution gradient program with mixture of water, methanol and methyl tert-butyl ether to separate, at least, 7 dominant pigments within 25 min. (Figure 5). The detailed identification of pigments, based on the chromatographic, spectrophotometric and mass properties, is summarized in Table 1. Chlorophyll a and chlorophyll b, α- and β-carotenes and violaxanthin are found to be the main chlorophylls and carotenoids, respectively, while the presence of lutein and zeaxanthin in this chloroplast is in low amount.
\nUFLC chromatogram of pigment extract from chloroplast of Pleomele angustifolia detected at 430 nm. The UFLC separation condition was as follows: Pigment separation was performed using UFLC equipped with PDA (Shimadzu) on C30 column (150 × 4.6 mm I.D; YMC) with a gradient elution program of water, methanol and methyl tert-butyl ether (MTBE) at the flow rate of 1 mL/min at 30°C.
Peak No | \ntR [min] | \nλmaxs [nm] | \nMolecular ion | \nFragment ions [m/z] | \nIdentification | \n|||
---|---|---|---|---|---|---|---|---|
HPLC eluent | \nHexane | \nEthanol | \nAcetone | \nspecies [m/z] | \n||||
1 | \n7.3 | \n412,436,464 | \n— | \n— | \n— | \n— | \n— | \nViolaxanthin | \n
2 | \n12.8 | \n470,601,650 | \n451,595,642 | \n465,601,649 | \n458,596,646 | \n907.7 [M]+ | \n881.7 [M – COH]+ 855.7 [M – COH – Mg]+ | \nChlorophyll b | \n
3 | \n13.4 | \n422,445,472 | \n422,444,473 | \n−,446,474 | \n−,448,476 | \n568.4 [M]+ | \n551.4 [M – OH]+ 476.4 [M – 92]+ 430.3 [M – 138]+ | \nLutein | \n
4 | \n15.3 | \n−,451,477 | \n425,449,478 | \n425,451, 478 | \n428,454,481 | \n568.6 [M]+ | \n476.4 [M – 92]+ | \nZeaxanthin | \n
5 | \n16.6 | \n431,618,664 | \n427,613,661 | \n430,616,664 | \n431,617,662 | \n893.5 [M]+ | \n871.5 [M – Mg]+ 615.2 [M – phytyl]+ | \nChlorophyll a | \n
6 | \n20.1 | \n421,446,473 | \n421,445,474 | \n421,446,476 | \n422,445,473 | \n536.6 [M]+ | \n445.4 [M + H – 92]+ | \nα-carotene | \n
7 | \n21.2 | \n–,452,478 | \n–,451,479 | \n–,453,480 | \n–,454,482 | \n536.6 [M]+ | \n444.5 [M – 92]+ | \nβ-carotene | \n
Chromatographic, spectrophotometric and mass properties of pigments separated from the chloroplast of Pleomele angustifolia.
Larger-scale separation of Chl a and b can be achieved by CC using Sepharose CL-6B as the stationary phase and a mixture of 2-propanol (IPA) and hexane as the mobile phase. Chl a could be eluted using 1.5% IPA in hexane and Chl b with 10% IPA in hexane [33]. To achieve a pure, free carotenoid, saponification step is sometimes necessary to eliminate contamination of lipids and chlorophylls. Moreover, carotenoid ester can be hydrolyzed to produce parent carotenoid by using this method [34]. CC is usually used for carotenoid isolation in high quantity of pigment extract. Generally, the purpose of CC is to separate mixtures into carotenoid fractions which are either having high purity to be processed to crystallization or low purity to be extensively separated with further chromatography, that is, HPLC [35].
\nSilica and alumina are frequently used as the absorbent in the CC with the normal phase elution to separate the distinct carotenoids; however, it is not easy to use this method to separate carotenoid isomers, that is, geometrical isomers, diastereoisomers, and so on. In this case HPLC/UFLC can be used to overcome the difficulty in the separation of carotenoids by CC. Turcsi et al. (2016) revealed that the polar carotenoids including optical isomers, and region and geometrical isomers as well as non-polar carotenes, could be well separated by HPLC on C18 and C30 columns, respectively [36]. High purity of isolated pigment can be achieved by HPLC and crystallization processes. UFLC analysis of the purified zeaxanthin shows that this carotenoid had a high purity of around 99.3% (Figure 2, left). All purified pigments have purity higher than 95% (Figure 6).
\nPurification of zeaxanthin: (a) chromatogram detected at 450 nm. Insert figure is UV–Vis spectrum measured by UFLC diode array detector in the eluent and (b) ESI-MS/MS spectrum identification. The conditions of UFLC and ESI-MS/MS analysis were as follows: UFLC analysis of the purified zeaxanthin was performed using UFLC equipped with PDA (Shimadzu) on C30 column (150 × 4.6 mm I.D; YMC) with a gradient elution program of water, methanol and MTBE at the flow rate of 1 mL/min at 30°C. The purified zeaxanthin was directly analyzed to LCMS 8030 (Shimadzu) with an isocratic elution of 0.1% formic acid (FA) in water (10%) and 0.1% FA in methanol (90%) at the flow rate of 0.3 mL/min. MS analysis was operated under the following conditions: (1) heat block temperature = 400°C; (2) desolvation line temperature = 250°C; (3) nebulizing N2 gas flow = 3 L/min; (4) drying N2 gas flow = 15 L/min; (5) interface voltage = 4.5 kV; (6) interface current = 0.1 μA; (7) mass range 400–700 m/z; (8) ionization mode = positive and negative.
Chromatographic, spectrophotometric and mass properties of pigment are minimum requirements for pigment identification [35]. These properties for all purified pigments are shown in the Table 1. In Figure 7 (right), absorption spectra of the purified chlorophyll a and the purified β-carotene in acetone have the same maximum absorption wavelength (λmax) and other spectral properties, such as the fine structure and spectrum shape, compared to these pigments in the references [37, 38]. Absorption spectrum of chlorophyll a in acetone shows typical Soret (431 nm), Qx (617 nm) and Qy (662 nm) bands, while two well-defined peaks in the absorption spectrum of β-carotene are found at 454 and 482 nm. This pigment analysis based on the results of spectrophotometer UV–Vis could support the advance pigment analysis using HPLC/UFLC equipped with photodiode array detection and coupled with the mass spectrometry. The LCMS technique has provided a power tool for pigment identification [39, 40]. Tentative identification for zeaxanthin peak separated by HPLC/UFLC analysis with PDA revealed that zeaxanthin has similar retention time (tR), maximum absorption wavelength (λmax) and the shape of absorption spectrum (data not shown) compared to the isolated zeaxanthin from corn which is a well-known source of zeaxanthin [41]. In addition the mass analysis provides the precursor and fragment ions at the specific m/z and characteristic fragmentation pattern for pigment identification. Mass spectrum of Chl a indicated the molecular ion [M]+ detected at m/z 893.6 and a fragment ion [M-Mg]+ at m/z 871.6 related to the loss of magnesium as the central metal of chlorophyll (Figure 1). This mass spectrum of Chl a agrees with the result that was reported [42].
\nPurification of Chl: (a) chromatogram detected at 660 nm. Insert figure is UV–Vis spectrum measured by UFLC diode array detector in the eluent and (b) ESI-MS/MS spectrum. The condition of UFLC and ESI-MS/MS analysis was as follows: UFLC analysis of the purified chlorophyll a was performed using HPLC equipped with PDA (Shimadzu) on C30 column (150 × 4.6 mm I.D; YMC) with a gradient elution program of water, methanol and MTBE at the flow rate of 1 mL/min at 30°C. The purified chlorophyll a was directly analyzed to LCMS 8030 (Shimadzu) with an isocratic elution of 0.1% formic acid (FA) in water (10%) and 0.1% FA in methanol (90%) at the flow rate of 0.3 mL/min. MS analysis was operated under the following conditions: (1) heat block temperature = 400°C; (2) desolvation line temperature = 250°C; (3) nebulizing N2 gas flow = 3 L/min; (4) drying N2 gas flow = 15 L/min; (5) interface voltage = 4.5 kV; (6) interface current = 0.1 μA; (7) mass range 400–1000 m/z; (8) ionization mode = positive and negative.
Chlorophyll and carotenoid are chloroplast pigments which are bound non-covalently to protein as pigment-protein complex and play a vital role in photosynthesis. Their functions include light harvesting, energy transfer, photochemical redox reaction, as well as photoprotection. The exact number and stoichiometry of these pigments in higher plants are varied, but their compositions include Chl a, Chl b, lutein, neoxanthin, violaxanthin, zeaxanthin and β-carotene. Liquid chromatography methods are well developed to separate and purify different types of pigments. Identification and characterization of pigments can be well observed by spectroscopy methods such as UV–Vis absorption, fluorescence and mass spectrometry.
\nTatas Hardo Panintingjati Brotosudarmo (THPB) acknowledges the competence research grant (No. 120/SP2H/LT/DRPM/IV/2017) from Kemenristekdikti for the financial support. We also acknowledge Chandra Ayu Siswanti who helped in preparation of chloroplast isolation, pigment isolation and UFLC separation works. We acknowledge Dr. Hendrik Octendy Lintang for supporting fluorescence measurements of photosystem II and I in chloroplast.
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