Examples of lanthanides and their concentrations in different plants and locations (according to Goecke et al. [3]).
\\n\\n
More than half of the publishers listed alongside IntechOpen (18 out of 30) are Social Science and Humanities publishers. IntechOpen is an exception to this as a leader in not only Open Access content but Open Access content across all scientific disciplines, including Physical Sciences, Engineering and Technology, Health Sciences, Life Science, and Social Sciences and Humanities.
\\n\\nOur breakdown of titles published demonstrates this with 47% PET, 31% HS, 18% LS, and 4% SSH books published.
\\n\\n“Even though ItechOpen has shown the potential of sci-tech books using an OA approach,” other publishers “have shown little interest in OA books.”
\\n\\nAdditionally, each book published by IntechOpen contains original content and research findings.
\\n\\nWe are honored to be among such prestigious publishers and we hope to continue to spearhead that growth in our quest to promote Open Access as a true pioneer in OA book publishing.
\\n\\n\\n\\n
\\n"}]',published:!0,mainMedia:null},components:[{type:"htmlEditorComponent",content:'
Simba Information has released its Open Access Book Publishing 2020 - 2024 report and has again identified IntechOpen as the world’s largest Open Access book publisher by title count.
\n\nSimba Information is a leading provider for market intelligence and forecasts in the media and publishing industry. The report, published every year, provides an overview and financial outlook for the global professional e-book publishing market.
\n\nIntechOpen, De Gruyter, and Frontiers are the largest OA book publishers by title count, with IntechOpen coming in at first place with 5,101 OA books published, a good 1,782 titles ahead of the nearest competitor.
\n\nSince the first Open Access Book Publishing report published in 2016, IntechOpen has held the top stop each year.
\n\n\n\nMore than half of the publishers listed alongside IntechOpen (18 out of 30) are Social Science and Humanities publishers. IntechOpen is an exception to this as a leader in not only Open Access content but Open Access content across all scientific disciplines, including Physical Sciences, Engineering and Technology, Health Sciences, Life Science, and Social Sciences and Humanities.
\n\nOur breakdown of titles published demonstrates this with 47% PET, 31% HS, 18% LS, and 4% SSH books published.
\n\n“Even though ItechOpen has shown the potential of sci-tech books using an OA approach,” other publishers “have shown little interest in OA books.”
\n\nAdditionally, each book published by IntechOpen contains original content and research findings.
\n\nWe are honored to be among such prestigious publishers and we hope to continue to spearhead that growth in our quest to promote Open Access as a true pioneer in OA book publishing.
\n\n\n\n
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Lanthanides play many roles in a number of different fields including chemistry, biology, and medicine [1]. They have also become indispensable in many modern technologies but the growing demand for these metals has also increased their release into the surrounding biosphere. Therefore, it is important to consider and address the impacts of increased lanthanides on the environment. The affinity of algae for these elements can pose a serious environmental threat or be a unique opportunity for the treatment of contaminated areas.
Lanthanides are considered nonessential elements that can induce both positive and negative physiological responses in the living organism. They are not essential for any known metabolic process, but under certain conditions, they may have a positive effect [2, 3]. Unlike heavy metals, whose toxicity has been extensively investigated, the effects of lanthanides have been neglected [4], particularly, their impacts on aquatic environments that are associated with the exploitation of lanthanides [5]. Water contamination by metals is a global problem, and metal recovery from wastewaters and industrial wastes is significant not only from an ecological point of view but also because of the sustainable availability of these materials [6].
This review aims to summarize our knowledge of positive and toxic effects of lanthanides on algae in order to better elucidate their biological roles. Various applications and methods of use, including the possibility of remediation and lanthanide recycling, are also summarized.
The presence of lanthanides (Pr, Nd, and Sm) was recorded for the first time in the red alga
Algae contain a diverse spectrum of lanthanides, regardless of size (micro or macroalgae), structural arrangements (unicellular, fibrous, and crustaceous), algal type (e.g., Chlorophyta, Rhodophyta, and Charophyta) as well as Cyanobacteria [8, 9, 10, 11]. These analyses show that seaweed lanthanide concentrations may be 10–20 times higher than those in terrestrial plants ([8], see Table 1) and more than 100 times higher than in sea water [10, 16].
Treea | Teab | Mossc | Potatod | Red algae | Brown algaf | Green algag | |
---|---|---|---|---|---|---|---|
Sc | nd | 0.085 | nd | nd | nd | nd | nd |
Y | nd | 0.360 | 0.127 | 0.011 | nd | nd | nd |
La | 0.280 | 0.600 | 0.266 | 0.017 | 0.362 | 0.032 | |
Ce | 0.370 | 1.000 | 0.493 | 0.038 | 0.943 | 0.076 | |
Pr | 0.091 | 0.120 | 0.056 | 0.007 | 0.049 | 0.008 | |
Nd | 0.155 | 0.440 | 0.402 | 0.015 | 0.191 | 0.039 | |
Sm | 0.031 | 0.085 | 0.036 | 0.008 | 0.034 | 0.009 | |
Eu | 0.004 | 0.018 | 0.009 | 0.001 | 0.008 | 0.028 | |
Gd | 0.024 | 0.093 | 0.037 | 0.007 | 0.044 | 0.012 | |
Tb | 0.017 | nd | 0.005 | 0.001 | 0.006 | 0.001 | |
Dy | 0.021 | 0.074 | 0.024 | 0.002 | 0.030 | 0.012 | |
Ho | 0.004 | 0.019 | 0.004 | 0.000 | 0.006 | 0.002 | |
Er | 0.006 | — | 0.013 | 0.002 | 0.015 | 0.008 | |
Tm | 0.001 | — | 0.001 | 0.000 | 0.002 | 0.001 | |
Yb | 0.008 | 0.044 | 0.011 | 0.001 | 0.008 | 0.007 | |
Lu | 0.019 | 0.007 | 0.001 | 0.000 | 0.001 | 0.001 | |
Examples of lanthanides and their concentrations in different plants and locations (according to Goecke et al. [3]).
Samples of
Certified reference material GBW07605 tea leaves, China [13].
Red alga
Brown alga
Green alga
The data correspond to mean values established in μg/g dry weight. In bold, the highest values of the series are highlighted.
Total lanthanides can range from 1 to 1.3 μg/g of algal biomass under laboratory conditions, and can be achieved easily, whereas under natural conditions (freshwater and sea water), the total amount of lanthanides ranges between 10−3 and 10−1 μg/g of algal biomass ([4, 17, 18, 19], and links therein).
There are only a few studies comparing lanthanides in different coexisting organisms, including algae. These studies indicate the relevance of lanthanides, particularly in microorganisms, and clear differences between coexisting groups of organisms (Table 2). Such a wide range of biotic concentrations of lanthanides can be generated by: (i) relative concentrations of elements in water; (ii) physical and metabolic processes specific to each type of algae (cell wall components, enzymes, proteins, etc.); and (iii) environmental factors specific to each area, e.g., temperature, light, pH, and nitrogen availability that can affect the two previous factors [22, 23, 24].
The concentration of lanthanides in the environment increases with changes in climatic conditions, groundwater action, and volcanic activity [25], but there are also significant anthropogenic sources of lanthanides in phosphoric mineral fertilizers, industrial waste waters, and mine extractions [4, 18, 26, 27, 28, 29]. Algae can serve as bioindicators because they can accumulate these elements in their cells (Table 1).
The probable biological effect of lanthanides is related to similarities between their ionic radii and coordination numbers with elements such as Ca, Mn, Mg, Fe, or Zn. Another aspect is their ability to form stable complexes with organic molecules [30]. Substitution of essential metal ions involves, for example, changes in enzyme activity, protein conformation, or polymerization. Also, changes in the use or allocation of ion channels affects specific membrane permeability and the cellular ion ratio.
Although lanthanides have been used for decades, particularly in China, as fertilizer in agriculture, their specific effects on plants and less so on algae, are not understood. Beneficial effects of lanthanides on growth and quality have been studied, mostly on crops [14, 31, 32] and domestic animals [14, 33, 34, 35]. Absorption, transmission, and metabolic conversion of nutrients were stimulated; metal deficiencies were overcome; and increases in metabolism via enzymatic activities were observed. Likewise, effects of lanthanides on photosynthesis or resistance to stress caused by drought, acid rain, and/or toxic metals (reviewed by [14, 32, 36, 37]) have been described. However, a specific cellular or molecular model for these observations has not been proposed and therefore mechanisms of action of lanthanide in plants or algae remain unclear [38].
One of the positive effects of lanthanides is connected with their ability to alleviate calcium deficiency because of Ln2+ and Ca2+ ions with high chemical similarities. These similarities, as well as the fact that lanthanides have higher valence values compared to calcium, resulted in Ln ions easily replacing Ca2+ and being able to bind with a higher affinity to multiple receptors, thus having various effects on metabolism depending on the effect of the replaced metal [31, 39, 40, 41, 42].
In the majority of experiments carried out with algae and lanthanides, attention was focused on algal (eventually cyanobacterial) growth properties without any effort to understand mechanism(s) of beneficial effects (Table 3). Thus, it is not clear whether the beneficial effects of lanthanides are due to the mitigation of nutrient deficiencies (such as Ca2+, Mg2+, or Mn2+), as previously found in plants [2, 48, 54, 55, 56], or to the fact that lanthanides are involved in some physiological reactions such as scavenging of oxygen-free radicals [30, 57, 58] or due to their ability to neutralize inhibitory effects of heavy metals [37].
Algae | Lanthanide | Positive effect | Negative effect | Reference |
---|---|---|---|---|
La3+ | 38.53–53 | >53.94 | [43] | |
* | LaCl3 | 30–40 | >40 | [44] |
Ce3 | 5–20 | — | [45] | |
La3+ | 5–20 | — | [45] | |
Ce3+ | 1.8 | 2.1 | [46] | |
* | 12 different Ln | — | 29.14 | [47] |
* | La3+ | <7.2 | >72 | [48] |
Dy3+ | 50–100 | 180–1000 | [49] | |
La | 7.28–87.4 | — | [50, 51] | |
Gd | 6.36–57.23 | — | [50, 51] | |
Yb | 5.78–17.34 | — | [50, 51] | |
La3+ | <7.2 | >72 | [48] | |
13 different Ln | — | 28–30 | [52] | |
Sc | — | 21.88 | [52] | |
Y | — | 43.21 | [52] |
Examples of studies testing the effect of lanthanides on growth, physiology, and survival of microalgae, specifying the concentrations at which positive, neutral, and negative effects were observed (values in μmol/L).
Algal divisions are characterized as Chlorophyta (C), Haptophyta (H), and Ochrophyta (O); Cyanobacteria (B) and Euglenophyta (E). If the algal species has a new name, it is referred to using the actual name and an asterisk (*); for names according to Algaebase, see Guiry et al. [53].
In a study on the effect of lanthanides in alleviating metal deficiency in algae, Li et al. [59] showed that La3+ at low concentrations were able to partly substitute for a Ca2+ deficiency in the green macroalga
Photosynthetic parameters expressed as maximum relative electron transport rates (rETRmax), and the maximal quantum yield (
The effects of single lanthanides and monazite on growth rate, lipid profile, and pigments in two biotechnologically interesting algae (
The use of lanthanides in agriculture and in aquatic cultures is gradually increasing although their impact on the environment has not been sufficiently verified. Lanthanides are not yet commercially available to increase the production of algal biomass despite the fact that their effects on economically interesting pigments and lipids are known. In the alga
The toxicity of lanthanides has been reported as low, but is dependent on their chemical form and processing, as reported by Hodge-Sterner’s classification system [62]. In soil and water, however, a surplus of lanthanides has a negative to toxic effect on human beings and animals [63]. Human exposure to lanthanides and effects on health are discussed by Pagano et al. [64]. The best studied effects on health are for Ce, La, and Gd, and the rest remain unclear [64]. The toxicity of lanthanides to various organisms is described in several reports [31, 42, 65], but maximum admissible concentrations, thresholds, and toxicity levels are poorly defined [66]. For each organism or species, the toxicity of different lanthanides differs, but the exact effects remain unknown [67, 68] (Table 3).
The ability of lanthanides to be involved in the metabolism of several basic elements has been considered as a possible cause of their toxicity [36]. Due to this phenomenon, differences in normal functions of several enzymes have been found, as demonstrated by work describing ATPase and pectate lyase [69, 70], ion channel blocking [71], or mineral transport [42, 72].
Although toxic effects of lanthanides have been reported for various microorganisms (Table 3), there is little evidence to generalize their effect on algae. Only a few orders of Charophyta [73], Chlorophyta [46, 48, 74], Dinophyta [75], Euglenophyta [49], Bacillariophyceae [76, 77] and Haptophyta [50], and Cyanobacteria [78, 79] have been studied. Most other algal studies, however, contained little or no data on the bioavailability of lanthanides. The relationship between lanthanide concentrations and stimulatory or inhibitory effects on the same algal species are therefore inconsistent. Moreover, many algal groups or species have not yet been tested for toxicity and no tests for macroalgae have been developed. The database on bioassays for algal toxicity is summarized in Guida et al. [80].
The transfer of lanthanides is expected through the food chain, as algae are primary producers [66, 81]. The toxicity of lanthanide on algae therefore needs to be addressed because any harmful effects may result in the transfer of negative effects to organisms at higher trophic levels [67, 82, 83].
Recent studies on the toxicity of lanthanides to algae describe the depletion of nutrients rather than toxicity itself [83, 84], see Section 7. In these works, it was suggested that lanthanides could capture some essential nutrients such as phosphates, resulting in an effect on growth (death by hunger). The relationship between lanthanides and phosphate was analyzed in detail in [85]. This important property should be examined in more detail because it could affect the bioavailability of these metals (EC50), changing the evaluation of their impact on the environment.
In recent decades, metal uptake by algal biomass has been studied with great interest. Uptake can be by passive binding, so-called “biosorption,” or an active process of “bioaccumulation,” where uptake or removal of elements is metabolically controlled [86, 87]. Some metals belong to the group of essential micronutrients, being important for growth and development of plant cells, and are involved in active metabolism [88]. Bioaccumulation of chemical compounds depends on rates of uptake and metabolism, and on the ability of the organism to degrade or store compounds. In essence, the process of accumulation of elements in algal cells is very complicated and depends on the properties of the species (type, size, form, and state of development), the element (charge, chemical form, and concentration), and the medium (pH, type, and concentration of metal salts or presence of complexing agents) [89]. As can be seen in Table 4, accumulation, biosorption, and desorption of lanthanides occurs in micro- and macroalgae, including brown, green, and red algae, algal flagellates, and also cyanobacteria. The potential for biosorption of cerium ions by cyanobacteria
Algae | Lanthanide | Reference |
---|---|---|
* | Ce | [90] |
14 different Ln, Y | [91] | |
Ce | [90] | |
Ce, La | [19] | |
La | [92] | |
* | Ce | [90] |
* | Ce, La | [19] |
Nd | [93] | |
Ce, Nd | [94] | |
Ce, La | [90] | |
Ce, La | [90] | |
Ce | [90] | |
* | Ce | [90] |
Eu, La, Yb | [95] | |
Eu, La | [96] | |
Eu, Gd, La, Nd, Pr, Sm | [1, 97] | |
Ce, La | [19] | |
Ce | [90] | |
Ce, Eu, La, Yb | [98] | |
14 different Ln, Y | [99] |
Studies on algal accumulation, biosorption and/or desorption of lanthanides.
Algal divisions Chlorophyta (C), Ochrophyta (O), and Rhodophyta (R), and Cyanobacteria (B), and the protist classes Dinophyceae (D) and Euglenophyceae (E) are specified. If microalgae were utilized, they are annotated with an (m). If an algal species has a new name, it is referred to with the actual name and an asterisk (*); names are according to Algaebase, see Guiry et al. [53].
Precise data about mechanisms of entry for lanthanides into algae and their accumulation are sparse. Even in higher plants, which are much more researched, cell processes responsible for lanthanide intake have only recently been described [38]. Several studies have shown that lanthanides concentrate in chloroplasts [93, 94, 106, 107, 108]. It was demonstrated that selective deposition of individual lanthanides in chloroplasts or the cytoplasm occurs in the green alga
Intracellular localization of different lanthanides in
However, many questions regarding the transfer and accumulation of lanthanides remain unanswered. For example, mechanisms of transport through the complex cell wall of algae or cyanobacteria, and whether they are stored in some specific structures or just loosely in the cytoplasm are unclear. Research into resistant strains or natural hyper-accumulators might bring some answers.
In biological systems, lanthanides are applied for different purposes such as growth promoters, fertilizers, water bloom killers, or as detection tools (bioindicators, tracers, and markers). Lanthanides have been proposed as growth stimulators for various animals such as pigs and other livestock [110]. Algae were also used as a feed additive to improve the condition of domestic animals [111]. Lanthanide-rich algae are a potential alternative to food supplements or functional foods. However, only one study on young abalones was performed to demonstrate that lanthanide-enriched algal biomass was an effective growth promoter [82]. Therefore, it would be important to increase the number of studies, to obtain relevant data on the effects of lanthanide transmission and to assess the risk of human exposure through food derived from animals [35].
Many microorganisms, including blue-green algae (e.g.,
The unique chemical features of lanthanides make them ideal tracers for geochemical processes in nature [9]. They represent alternative, nonradioactive, highly detectable labels. They were used, for example, to confirm the impact of cyanobacterial mats on deep waters outside French Polynesia, providing evidence for an end-ascending flow [114]. They enable scientists to follow oceanic cycles, petrogenesis, the chemical evolution of the Earth [16, 29], or palaeo-environmental conditions [115, 116, 117, 118]. Lanthanides can also serve as anthropogenic activity indicators [27]. Because of their particular affinity to algae, the lanthanide profile may be a useful indicator for exploring the ecology of marine environments [10] and can also be used to monitor sources of pollution from natural events such as volcanic activity [25]. In combination with macroalgal sampling, the lanthanide profile may help to characterize coastal water quality and pollution [22, 23, 27].
Lanthanides have been used for their inert nature as detection agents in various experiments, for example, in studies of the rate of passage and digestibility of nutrients in humans and animals [119, 120, 121]. Lanthanide oxides have been used as markers in sea cucumber (
In the development of new, sensitive detection methods, active chelates of lanthanides have been obtained and tested. They are used in sensitive immunoassays to suppress the background [122] or as very sensitive fluorescence probes [123]. An example of their use is the labeling of the cyanotoxin microcystin [124, 125].
In countries with sufficient sources of lanthanides (mainly China), these elements are used as fertilizers to increase agricultural production. With increasing consumption, waste with varying contents of different lanthanides has increased significantly and rapidly. The most important of these are magnets (neodymium), metal alloys (europium and yttrium), batteries, glass, and catalysts (cerium and lanthanum) [126]. Other important sources of lanthanide waste are phosphate mineral fertilizers, industrial wastewater, sewage sludge, mining processes, or wastes from industrial aluminum production [4, 18, 26, 27, 28, 29]. Lanthanides present in ecosystems from agricultural production can thus penetrate into the groundwater and migrate to rivers and lakes [58] or to the sea [127]. Some studies on ecological effects and potential threats due to the bioaccumulation of lanthanides have been described, but they are not long-term enough to draw any general conclusions [128, 129]. Relevant regulations or standards concerning doses and threshold values for the presence of lanthanides in the environment have also not been established [38]. In China, lanthanides are cited as the main source of environmental contamination [130]. They are also considered to be emerging pollutants outside of China, requiring the specification of threshold values for concentrations and emissions of lanthanides in the environment [64, 131]. Removing these lanthanide contaminants is therefore a very important requirement in order to reduce the ever-increasing environmental burden on the aquatic environment.
In addition to this very important requirement for remediation, the need for recycling of lanthanides from any (not only liquid) industrial production waste becomes even more acute. One reason is the risk of reduced availability of resources (China owns more than 95% of natural sources) or their relatively rapid depletion from other sources. Replacement of lanthanides with alternate substances in industrial applications is currently not possible [132, 133]. Due to their unique chemical and physical properties and their extensive applications in industrial products, the importance and demand for these elements is constantly increasing [131, 134]. The economic impact of an emerging lanthanide shortage increases the urgency for efficiently using renewable energy sources from the ever increasing number of different types of waste products worldwide. At present, research is focused on the progressive and cost-efficient recycling of lanthanides for industrial processes [4, 95, 102, 135, 136], which would reduce risks associated with inaccessibility or depletion of natural resources while minimizing environmental problems associated with their extraction and processing [137].
One of the most widespread lanthanide-containing wastes is electrical and electronic equipment, including lighting equipment, computers, or photovoltaic panels. This waste is a growing threat to the world’s environment, and lanthanide recovery is therefore becoming economically attractive. The main sources for recycling are luminophores, powder mixtures obtained from electronic waste and containing high concentrations of lanthanides. Luminophores are obtained from television screens or monitors, as well as energy-saving bulbs and lamps, where they are used to convert cathodic tube radiation or ultraviolet electric discharge into mercury vapor and visible light. These luminophores occur as a powder attached to the inner surfaces of mesh or tubes. The glass parts of these waste networks, monitors, screens, and light bulbs can be easily recycled, but luminophore layers must be removed because the luminescent compounds would reduce the quality of recycled glass. The luminophores as waste represent a toxicity problem but, on the other hand, are a concentrated source of various lanthanides, either in the form of dry powder or wet mud [138].
Lanthanides from waste sources can be recycled by chemical separation from solutions (e.g., chemical precipitation, electrochemically, membrane division, reverse osmosis, etc.). These methods are comparatively costly and, moreover, are often a source of other nonorganic wastes [139]. Methods such as pyrometry and hydrometallurgy for the extraction of lanthanides from ores have significant negative impacts on the environment and involve high costs [126]. The other serious disadvantage is the dependence on a single and limited source and possibly the depletion of other natural resources [126, 140, 141]. These traditional physicochemical processes are expensive or even inefficient for the treatment of sewage containing low concentrations of metal ions [142]. A by-product of conventional methods is the associated large volume of contaminated water, high temperatures and a high consumption of chemical compounds [143, 144]. Researchers are therefore looking for low-cost approaches and at the same time environmentally friendly technologies.
As a biotechnological approach, biosorption is considered to be a more efficient and cheaper alternative to conventional chemical methods of recycling lanthanides [133, 145, 146]. Various different organic residues of animal or plant origin, including resin, activated charcoal, or biomass of various organisms (algae, fungi, and bacteria), have been shown to adsorb different lanthanides and have been tested as biosorbents [95, 98, 132, 147]. The development of effective biological methods for lanthanide regeneration from these materials was proven in the aerobic, genetically modified bacterium,
Methods for the recycling of lanthanides via living cells offer an alternative, which does not have the disadvantages of chemical and adsorption approaches. Accumulation of lanthanides from the environment is cost-effective and does not produce any substantial secondary waste. In addition, it is a great advantage that it can also be effective in water containing very low lanthanide concentrations, which is problematic in other approaches.
Waste solutions containing lanthanides often have high acidity. Thus, the discovery that the sulfothermophilic red alga
Up to now, only one paper has been published demonstrating the high potential of seaweed (in this case, brown algae
Algae are very important organisms in terms of ecology, being at the very beginning of the food chain. Their relationships with metals therefore affects other living organisms. Their ability to accumulate lanthanides may have an impact on the surrounding environment, representing both a threat and an opportunity, with the potential for further study and use. As bioaccumulation abilities and beneficial or toxic effects of lanthanides differ in individual algal strains, it is difficult to predict specific ecological hazards. Algae in combination with lanthanides offer a wide variety of applications. They can be used as bioindicators, fertilizers, toxin detectors, or for phytoremediation and recycling. Therefore, understanding the relationships between algae and lanthanides is very important. Once we understand the molecular mechanisms of their effects, we will have greater opportunities for their use.
We thank Prof. John Brooker for critical reading and language corrections of the manuscript. This work was supported by the National Program of Sustainability I, ID: LO1416.
Chlorophyll 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)
Plant 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
Chemical structure of Chl
(a) UV–Vis absorption of Chl
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
In 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) 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 (
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) 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
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
Unlike 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
The 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
Chlorophyll 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
After 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
UFLC chromatogram of pigment extract from chloroplast of
Peak No | \nt | \nλmaxs [nm] | \nMolecular ion | \nFragment ions [ | \nIdentification | \n|||
---|---|---|---|---|---|---|---|---|
HPLC eluent | \nHexane | \nEthanol | \nAcetone | \nspecies [ | \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 | \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 | \n
6 | \n20.1 | \n421,446,473 | \n421,445,474 | \n421,446,476 | \n422,445,473 | \n536.6 [M]+ | \n445.4 [M + H – 92]+ | \n|
7 | \n21.2 | \n–,452,478 | \n–,451,479 | \n–,453,480 | \n–,454,482 | \n536.6 [M]+ | \n444.5 [M – 92]+ | \n
Chromatographic, spectrophotometric and mass properties of pigments separated from the chloroplast of
Larger-scale separation of Chl
Silica 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
Purification 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
Tatas 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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