ISSN 1021-4437, Russian Journal of Plant Physiology, 2006, Vol. 53, No. 2, pp. 257–277. © MAIK “Nauka /Interperiodica” (Russia), 2006. Original Russian Text © I.V. Seregin, A.D. Kozhevnikova, 2006, published in Fiziologiya Rastenii, 2006, Vol. 53, No. 2, pp. 285–308. REVIEWS Physiological Role of Nickel and Its Toxic Effects on Higher Plants I. V. Seregin and A. D. Kozhevnikova Timiryazev Institute of Plant Physiology, Russian Academy of Sciences, Botanicheskaya ul. 35, Moscow, 127276 Russia; fax: 7 (095) 977-8018; e-mail: ivanov@ippras.ru Received April 18, 2005 Abstract—The focus of the review is on the specific aspects of nickel effect on plants as compared to other heavy metals; their specificity is derived from different physical and chemical properties. The various facets of the physiological role of nickel and its toxic activity in higher plants, its intracellular partition and transport in plant tissues and organ are discussed. The putative mechanisms of nickel hyperaccumulation are considered in several representatives of angiosperm plant families. The existing evidence was used to outline the metabolic changes in plants affected by nickel. The comparison with other heavy metals is used to disclose the general mechanisms that disturb plant mineral nutrition, water regime, photosynthesis, and morphogenesis as well as the common cell responses aimed at detoxification of heavy metals. The numerous nonspecific effects of heavy metals depend on their direct and indirect action; in addition, some effects of nickel are specific. To illustrate, high Ni content in endoderm and pericycle cells blocks cell divisions in the pericycle and results in the inhibition of root branching. DOI: 10.1134/S1021443706020178 Key words: higher plants - mechanisms of activity - nickel - stress - tolerance INTRODUCTION Nickel holds a special place among the heavy metals. Unlike Cd, Pb, Hg, Ag, and several other metals that are not the components of plant enzymes, Ni is a constituent of urease, and small quantities of Ni (0.01 to 5 µg/g dry wt) are essential for some plant species. On the other hand, Ni is not as important for plant metabolism as Zn and Cu. However, same as with other heavy metals, high Ni concentrations may turn toxic to plants. The analysis of published evidence on Ni toxicity towards plants shows that, in addition to general toxicity displayed by all heavy metals, Ni manifests the specific characteristics due to its characteristic physical and chemical properties. The toxicity of heavy metals may depend on their binding to various ligands; among such ligands in the biological systems, carboxylate ion, imidazole, sulfhydryl group, and aliphatic amine are the most important. To illustrate, heavy metal binding to various functional groups of proteins, primarily SHgroups, would modify protein conformation and result in the loss of activity in many enzymes comprising SHgroups in their activity centers. Besides, metal binding to SH-groups of the physiologically active compounds of low molecular weight would also interfere with cell metabolism. Thus, binding to SH-groups alone would already give rise to diverse metabolic disturbance. Heavy metals differ in their affinity for oxygen-, nitrogen- and sulfur-containing ligands, and these differences depend on the physical and chemical proper- ties of heavy metal ions. The metal cations are classified into hard and soft Lewis acids [1]. According to this classification, the ions of hard acids, such as Li+, Sr2+, etc., would rather interact with hard bases, whereas the ions of soft acids, such as Pb2+, Hg2+, Ag+, etc., with soft bases. Cd2+, Cu2+, Ni2+, and several other ions come into an intermediate class, with Ni2+ shifting towards hard and Cd2+, soft class. In the biological chemistry, there are four major classes of donor atoms in ligands. Among these, oxygen and aliphatic nitrogen are classified as hard, aromatic nitrogen as intermediate, and sulfur as soft. It follows that Ni would rather bind to aromatic nitrogen, and Cd2+, Hg2+, and Ag+, to SH-groups. The rate of ligand exchange within and beyond the coordination sphere of the particular metal ion is an important parameter of its reactions. For example, Ni2+ and Zn2+ practically do not differ in several physical and chemical properties, such as the size of the ion. However, Zn2+ is a constituent of numerous enzymes, whereas Ni2+ is found in few plant enzymes, probably because in the former case the rate of ligand exchange is higher by three orders of magnitude. Thus, diverse effects of the metals of similar properties apparently depend on the different rates of ligand exchange, and their physiological properties are immediately derived from their physical and chemical parameters. Because of the competition between various metals in the course of their uptake by roots, some metals are absorbed in insufficient quantities, whereas the uptake 257 258 SEREGIN, KOZHEVNIKOVA of other metals is excessive. Such situation would indirectly predetermine the effects of heavy metals on the various facets of metabolism, such as photosynthesis, respiration, etc. It also seems important to compare the mechanisms of heavy metal accumulation, transport, toxicity, and detoxification in susceptible and tolerant plant species and populations. To this end, it was crucial to review the characteristic aspects of Ni2+ transport and distribution and Ni2+ effects on various physiological processes. In this aspect, Ni2+ is of particular interest due to numerous hyperaccumulators of nickel already discerned in various plant families. PHYSIOLOGICAL ROLE OF NICKEL IN HIGHER PLANTS Currently a metal is considered to be an essential nutrient in the cases when plants cannot complete their life cycle in its absence and it cannot be substituted with any other nutrient [2, 3]. In 1987, Ni was first established as a nutrient essential for completion of plant life cycle. Ni deficiency decreased the capacity of barley to develop viable seeds because of hindered embryo growth. The embryonic root developed poorly or even stayed undeveloped; in addition, several anomalies were reported in endosperm development together with declined dehydrogenase activities. The critical Ni concentration in barley tissues that reduced the yield by 15% was 90 ± 10 ng/g dry wt [4]. In the middle of 1970s, Ni2+ was shown to be a constituent of urease, a metalloenzyme from Canavalia ensiformis seeds, engaged in urea hydrolysis [5, 6]. Later Ni2+ was also found in urease from other plant species [7, 8]. Presently three basic pathways of urea biosynthesis are known to function in higher plants [9–11]. First, it is the terminal step of the ornithine cycle catalyzed by the mitochondrial arginase; this step involves arginine hydrolysis to ornithine and urea. Second, it is purine catabolism leading to formation of ureides, such as allantoin and allantoinic acid; the latter is hydrolyzed to glyoxylate and urea in the reaction catalyzed by allantoicase. This pathway is characteristic of the Fabaceae plants, which accumulate ureides as the major reserve form for translocating nitrogen from roots into shoots. In addition, arginine and ureides are often the reserve forms of nitrogen in the seeds. Finally, an additional pathway of urea synthesis from a nonprotein amino acid canavanine was found in Canavalia and soybean. Urea derived from these reactions is hydrolyzed by urease, and the liberated ammonium participates in various anabolic processes, particularly in glutamine synthesis from glutamate run by glutamine synthetase, a crucial reaction in the glutamine synthetase/glutamate synthetase pathway of ammonium assimilation. In the experiments with rye, rapeseed, zucchini, and sunflower, Ni increased the glutamine content and also, in two latter plant species, the content of the ornithine cycle acids [12, 13]. There are two types of ureases: a tissue-specific enzyme found in the vegetative tissues of most plant species and an embryonic enzyme, a characteristic seed protein in soybean, Arabidopsis, Canavalia, etc. In soybean, tissue and embryonic ureases are encoded by different structural genes Eu1 and Eu4, respectively [7]. The amino acid sequences of all known ureases are of considerable homology, presuming similar tertiary structures and mechanisms of activity [8]. Plant and fungal ureases are homooligomeric proteins, whereas bacterial ureases are multimers comprising two or three different subunits. Thus urease from Canavalia is a homotrimer which can form a homohexamer [14]. The polypeptide chain of this urease comprises 840 aminoacid residues, with 13 out of 25 histidines concentrated in the region between residues 479 and 607. Apparently here, the active center of the enzyme containing Cys592 residue bound to Ni ions and essential for enzyme activity is located [15]. Thus, both SH- and Nligands are necessary to bind Ni2+ in the urease active center because of high Ni2+ affinity for intermediate and soft bases. One urease subunit comprises two Ni2+ ions crucial for the catalytic process: they bind the substrate; in addition, these ions bind urease inhibitors. Thus, urease of Klebsiella aerogenes consists of three subunits each containing two Ni2+ ions in their active centers. One of these Ni ions is bound to two histidine residues (His246 and His272), and the second, to amino-acid residues His134, His136, and Asp360. In addition, Lys217 helps secure two Ni ions [8]. In spite of the crucial role of nickel in urease formation, this nutrient does not induce urease synthesis [16]. To secure Ni2+ in the urease active center, several auxiliary proteins are engaged as urease-specific chaperons. Similar to ureases from various organisms, these chaperons are of considerable structural homology [8]. In soybean, Ni2+ attachment to the enzyme depends on the activities of two genes, Eu2 and Eu3 encoding the auxiliary proteins that activate urease. The mutations in these genes resulted in the loss of urease activity; however, these mutations practically did not affect enzyme content and Ni uptake and translocation [7]. The Eu3 gene encodes a 32-kD protein, which interacts with the product of Eu2 in the course of the embryonic urease activation. When this process was blocked with the antibodies against the protein Eu3 accumulating in developing embryos, the urease content did not increase because the enzyme was instable when devoid of Ni2+ in its active center [8]. This evidence shows that a sufficient quantity of available Ni is essential for urease activation. A series of experiments with various plant species and growth media was run to demonstrate that Ni and urease are essential for plant vital functions. The deficiency in Ni content in the medium and the low activity of urease RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL resulting from such deficiency upset nitrogen metabolism and led to the accumulation of toxic urea levels in shoots; phenotypically such process was manifested as necroses in leaf tips or chlorosis in older leaves [2, 9, 10, 12, 13, 17, 18]. Similar necrosis of leaf tips was observed following soil dressing with high rates of urea. It was urea and not ammonia that caused the necroses because the addition of a urease inhibitor augmented the necrosis [19]. Leaf injury was especially manifest in plant species that were capable to develop the symbiosis with nitrogen-fixing bacteria: in such species, nodule development on the roots lagged behind by two or three days [2]. The addition of Ni salts at low concentrations to the nutrient solution alleviated these symptoms; Ni could not be substituted with other nutrients, such as Al, Cd, Sn, V, Cr, and Pb [20]. Urease plays an important role in seed germination. The addition of urease inhibitors retarded germination, apparently because urea accumulated to toxic levels [11]. Low Ni concentrations were shown to promote seed germination in rice [21], wheat, timothy grass, pea, bean, soybean, and lupine (cited after [9]). In addition to urease, low concentrations of Ni were reported to activate several more enzymes [3, 9, 10]; however, the mechanism of this phenomenon stays unknown. In some legumes, small amounts of Ni are essential for root nodule growth and hydrogenase activation. The efficiency of nitrogen fixation immediately depends on hydrogenase activity because the oxidation of hydrogen by the latter provides ATP required for N reduction to ammonia. Ni deficiency was shown to lower down hydrogenase activity in the nodules. On the contrary, when soybean plants grown in soil culture were irrigated, once in two weeks, with the nutrient solution containing 1mM NiCl2, at day 52 the hydrogenase activity of Rhizobium japonicum nodules exceeded that of the control plants by about 45%, although the promoting effect disappeared by day 100, apparently as a result of increasing Ni toxicity [17]. The fact that several enzyme activities depend on the presence of Ni ion can explain the promotional effects of low Ni concentrations on plant growth and development in such species as zucchini, oilseed rape, cotton, sweet pepper, tomato, potato, and Chinese hemp [9, 12, 13, 18]. Thus, spraying cotton plants with nickel sulfate solution (234.8 mg/kg) increased the numbers of buds and flowers, the rate of boll formation, and seed oil content (by 4.6%) [3]. Thus, we demonstrated that higher plants need Ni, and therefore Ni is classified among the essential ultramicronutrients. Ni UPTAKE BY PLANTS Nickel is delivered into the environment by several pathways: (1) as factory waste of high-temperature technologies of ferrous and nonferrous metallurgy, cement clinker production, and burning liquid and solid RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 259 fuels; (2) field irrigation with water high in heavy metal content and transfer of sewage residue into soil; (3) transfer of heavy metals from mine tailings and metallurgical factories by water and air flows; (4) steady application of high rates of organic and mineral fertilizers and pesticides contaminated with heavy metals [22]. On the average, the total Ni content in soil varies from 2 to 750 mg/kg soil, with the maximum content reported in the serpentine soils [1]. The major nickel ores are garnierite [(Ni, Mg)6Si4O10(OH)2] and penlandite [(Ni, Fe)9S8] [23]. The major way of Ni uptake from soil is its absorption by plant roots. Some of the soil-accumulated metal becomes bound to its organic components and becomes inaccessible to plants, whereas metal ions can enter the roots. Currently few papers describe the mechanism and kinetics of Ni2+ absorption by plants. Plant absorption of Ni2+, same as of other metals, may proceed due to passive diffusion and active transport. To elucidate the role of metabolic processes in Ni2+ absorption, the rates of Ni2+ translocation were compared at various temperatures and in relation to the aeration of nutrient solutions. At 23°C, Ni2+ uptake by Avena sativa roots directly depended on the incubation period. Low temperature, such as 2°C, considerably lowered Ni2+ absorption from the nutrient solution. The relationship between Ni2+ absorption and temperature was described by an S-like curve, with the maximum between 23 and 30°C. Both the addition of 20 µM 2,4dinitrophenol to the nutrient solution and the anaerobic conditions of plant growth inhibited Ni uptake by 91 and 86%, respectively. These data presume that the metabolically active uptake considerably exceeds the passive entry of Ni2+ ions [24]. The ratio between the inputs of active and passive components depends on Ni2+ concentration in the nutrient solution. The former is more important at low Ni2+ concentrations (below 34 µM), and at higher concentrations the role of passive transport mechanism increased due to Ni2+ toxic effect [25]. The major environmental factors that affect metal uptake by plants are soil acidity, its cation exchange capacity, the contents of organic substance and lime, moisture potential, granulometric composition, and the concentrations of macro- and micronutrients [1, 3, 24, 26–28]. The effects of these factors on the uptake of many heavy metals are mostly nonspecific. The accessibility of Ni usually declines at higher pH values of the soil solution due to the formation of the low-soluble complexes [1, 25]. Besides, in soil Ni is strongly absorbed by magnesium and calcium oxides in the wide pH range and by iron oxide at pH above 5.5 [1]. As a result, the surface of roots and soil particles would be coated with the film of insoluble iron (III) compounds, which can bind Ni2+ and make it inaccessible to plants [29]. Thus, in several populations of Alyssum murale and A. corsicum, the shoot content of Ni was No. 2 2006 260 SEREGIN, KOZHEVNIKOVA shown to increase at the elevated pH values of soil solution [28]. Higher Ni mobility was also reported in the soils with lower humus content, lighter granulometric composition, and higher moisture content [27]. When absorbed by roots, Ni2+ ions may compete with other metal ions. To illustrate, the antagonistic relations were described for Ni2+ and Zn2+ in such hyperaccumulator species as Thlaspi montanum, T. caerulescens, Streptanthus poligaloides, and Dichapetalum gelonioides, with addition of Ni2+–Fe2+ antagonism in T. montanum; similar antagonistic relations were reported in Alyssum bertolonii between Ni2+, Zn2+, and Co2+ and in plant species that exclude heavy metals, such as Glycine max and Hordeum vulgare, Ni2+ uptake declined in the presence of Cu2+ and Zn2+ (cited after [30]). Calcium ions most strongly affect the uptake of heavy metals from the soil solution; however, these effects considerably varied. Thus, Gabbrielli and Pandolfini [31] demonstrated that Ca2+ lowered Ni2+ absorption by A. bertolonii, an endemic plant of serpentine soils, and promoted Ni2+ absorption in Berkheya coddii [30]. Apparently, various plant species employ different molecular mechanisms for heavy metal absorption. Some researchers suggested that Ni2+ enters plant cells via activated calcium channels [30, 32], whereas in grasses, the uptake of Ni2+ and some other heavy metals may involve phytosiderophores; in this context, the latter were called phytometallophores [33]. No specific Ni2+ transporters have been as yet described [34, 35]; nevertheless, this process can employ various nonspecific transporters. As a whole, the inhibitory effect of metal ions on Ni2+ absorption and translocation from roots into shoots decreased in the following order: Fe3+ > Co2+ > Ca2+ > + Mg2+ > NH 4 > K+ > Na+ [25]. Such range is in good accordance with the evidence for other heavy metals [36] and apparently signifies their nonspecific effect on Ni2+ uptake. In addition to absorption by the roots, Ni can enter the plants via the leaves. When 63Ni was applied to the leaves of Helianthus annuus, 37% of the total amount was translocated into other plant organs (cited after [25]). Similar patterns were observed when oat, soybean, tomato, and eggplant leaves were sprayed with solutions of Ni salts (cited after [3]). Thus, the specific mechanisms of Ni+ uptake have not been as yet disclosed. It is not clear whether the hyperaccumulator species acquired particular absorption mechanisms selective towards Ni2+ and the ability to increase its accessibility. It is not known whether the tolerance of particular plant species relies on the lowered Ni2+ uptake or, quite the reverse, depends mostly on the characteristic patterns of Ni2+ translocation and distribution and binding Ni into insoluble complexes. Below we will consider numerous and widely conflict- ing data concerning Ni transport and allocation in plant organs. NICKEL DISTRIBUTION IN PLANT ORGANS: THE MECHANISMS AND ROLE OF HYPERACCUMULATION Plant species vary in their capacity to accumulate heavy metals. High accumulation of heavy metals and the ratio of metals in various organs of diverse plant species largely depend on plant morphophysiological characteristics. The current classification divides all plant species into three groups: (1) the accumulators that store metals mainly in the shoots under high and low metal concentration in soils; (2) the indicators, with plant metal concentrations reflecting the metal content in the environment; and (3) the excluders, with restricted transfer of heavy metals into the shoots whatever high are metal concentrations in the environment and the roots [37, 38]. Among the plants accumulating Ni, there is a discrete group of the hyperaccumulators that accumulate metals in the shoots to the level of over 1000 mg/kg dry wt [39]. Presently about 300 such species have been described; they mostly belong to the families Asteraceae (27), Brassicaceae (82), Buxaceae (17), Euphorbiaceae (83), Flacourtiaceae (19), Rubiaceae (12), and Violaceae (9) and grow on serpentine soils in the tropical and subtropical zones of Cuba, New Caledonia, Indonesia, Philippines, Brazil, Australia (Queensland), the South Africa (Zimbabwe), and the Mediterranean region [40] (Table 1). Thus, the Cuban flora comprises 130 hyperaccumulators mostly representing the genera Buxus, Phyllanthus, Leucocroton, Euphorbia, Pentacalia, Senecio, Psychotria, Ouratea, and Tetralix [41]. The number of species that hyperaccumulate other heavy metals is considerably lower, and elucidating the cause of this phenomenon is an inviting problem. There are contradictory hypotheses as to the mechanism of hyperaccumulation. Thus, Boyd and Martens [30] presume that the hyperaccumulators possess the most efficient system of ion absorption with yet unknown functions. There are also several hypotheses explaining the role of hyperaccumulation in plant vital functions. Within the removal hypotheses, the hyperaccumulation is seen as a mechanism of tolerating high levels of metal in the environment: in this case, the absorbed metal is transferred into the compartments of low physiological activity or into plant organs to be shed in future [30, 42]. Indeed, the old leaves of the hyperaccumulator species Psychotria douarrei stored considerably more Ni than the young leaves [43]. The drought resistance hypothesis claims that Ni accumulation in plant tissues would enhance plant tolerance to moisture deficit by reducing the cuticular transpiration [30, 42, 44]. According to the hypothesis of elementary allelopathy, shedding plant organs with high heavy metal content, such as leaves, would enrich soil surface RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL 261 Table 1. Plants hyperaccumulating nickel and their capacity to amass this heavy metal Family, species Maximum concentration, mg/kg dry wt Acantaceae Blepharis acuminata Justicia lanstyakii Lophostachys villosa Phidiasia lindavii Ruellia geminiflora Anacardiaceae Rhus wildii Asteraceae Berkheya coddii Dichoma niccolifera Gochnatia crassifolia Koanophyllon grandiceps Leucanthemopsis alpina Senecio pauperculus Shafera platyphylla Solidago hispida Brassicaceae Allysum (48 species) Cardamine resedifolia Cochlearia aucheri Peltaria emarginata Streptanthus polygaloides Thlaspi (23 species) Caryophyllaceae Minuartia laricifolia Minuartia verna Clusiaceae Garcinia bakeriana Dipterocarpaceae Shorea tenuiramulosa Euphorbiaceae Cleidion viellardii Euphorbia (3 species) Leucocroton (28 species) Phyllanthus (41 species) Savia (3 species) Fabaceae Pearsonia metallifera Trifolium pallescens Flacourtiaceae Casearia silvana Homalium (7 species) Xylosma (11 species) Family, species Meliaceae Walsura monophylla Myristiaceae Myristica laurifolia Myrtaceae Mosiera araneosa Ochnaceae Ouratea nitida Ouratea striata Oncothecaceae Oncotheca balansae Poaceae Trisetum distichophyllum Ranunculaceae Ranunculus glacialis Rubiaceae Phyllomelia coronata Psychotria clementis P. constivenia P. douarrei P. glomerata P. vanhermanii Sapotaceae Planchonella oxyedra Sebertia acuminata Saxifragaceae Saxifraga (3 species) Scrophulariaceae Esterhazya sp. Linaria alpina Stackhousiaceae Stackhousia tryonii Tiliaceae Tetralix brachypetalus Turneraceae Turnera subnuda Villoziaceae Villozia sp. Violaceae Agatea deplanchei Hybanthus (5 species) Rinorea bengalensis R. javanica 2000 2690 1890 1850 3330 1600 11660 1500 1120 6240 3200 1900 1890 1020 1280–29400 3270 17600 34400 14800 2000–31000 2710 1390 7440 1000 9900 4430–9340 2260–27240 1090–60170 2940–4890 15350 1990 1490 1160–14500 1000–3750 Note: Modified data from [40]. RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 Maximum concentration, mg/kg dry wt 7090 1100 5030 1700 8190 2500 1710 1260 25540 19090 38530 19900 10250 35720 19600 11700 2970–3840 1060 1990 21500 13610 6130 3080 2500 3000–17600 17500 2170 262 SEREGIN, KOZHEVNIKOVA with these metals and inhibit the growth of competing neighbor species that are not tolerant to the corresponding heavy metals [30, 42]. According to the most widely accepted defense hypothesis, the elevated concentrations of heavy metals, especially in the plant dermal tissues, would avert the penetration and propagation of pathogenic microorganisms. In addition, such hypothesis supports the view that the accumulator plant species are in advantage over non-accumulators because the former spend less nitrogen and carbon on organic substances, such as tannins and phenolics that protect plant tissues against the herbivores, mostly insects [30, 42, 43, 45, 46]. The defense hypothesis was supported by several experimental studies. Thus, the pathogenic strain of Xanthomonas campestris pv. campestris did not develop in Streptanthus polygaloides plants when they accumulated Ni, and the growth of the parasitic fungus Erysiphe polygoni and the necrotrophic fungus Alternaria brassicola was inhibited [47]. The larvae of Pieris rapae failed to pupate and died when fed with the leaves of S. polygaloides plants grown on the soil rich in Ni. When Ni or latex from another hyperaccumulator, Sebertia acuminata, was added to the feed of Drosophila melanogaster larvae, they starved, and the singular survivors did not develop normally [45]. Most plant species belong to the excluder group. Thus, Ni is predominantly accumulated in the roots of Zea mays [48, 49], Typha latifolia [29], Alyssum montanum, and Thlaspi arvense [50, 51], Glycine max [17], Avena sativa (at the tillering and booting stages) [52], Cyperus difformis, Chenopodium ambrosioides, and Digitaria sanguinolis [53], Triticum aestivum and Hordeum vulgare [3] (Table 2). However, when Ni accumulation exceeds a characteristic limit, plant mechanisms of detoxification that keep low the shoot metal concentrations become less efficient. As a result, the control over metal entry into the shoots is lost, and such plants perish [36, 38]. Nickel distribution in plants depends on their developmental stage. Thus, most Ni accumulation in the roots of Avena sativa was registered at the tillering and booting stages, while the maximum accumulation in the reproductive organs proceeded at the heading stage and at the stages of milky and full maturity [52]. This evidence leads to an important practical question: what is the mechanism that determines the capacity of the hyperaccumulator species to hoard Ni in the shoots and to avoid its toxic effects. Apparently, no sole mechanism exists for Ni accumulation in the shoots and no sole mechanism stands for the tolerance of these particular plant species towards the excess of Ni. As a whole, the process of hyperaccumulation seems to depend on the characteristic patterns of metal translocation and distribution, while their concomitant tolerance to the excess of metal relies on several detoxification mechanisms. Below we will consider these issues in more detail. NICKEL DISTRIBUTION IN PLANT TISSUES Infrequent papers on Ni tissue distribution deal mainly with its localization in the shoots of hyperaccumulator plant species as established from the data obtained using the histochemical (dimethylglyoxime) method [44, 45, 60, 61], the spectroscopic and atomic absorption spectrometry [62, 63], X-ray microanalysis, and scanning electron microscopy [60, 61]. Thus, in the seeds of the hyperaccumulator Thlaspi pindicum collected on serpentine soils, Ni was discovered mainly in seed coats, with the maximum accumulation around the micropile and in the epidermis and lower amount in the mesophyll cells of cotyledons [63]. This evidence matches the data that Ni is accumulated in the leaf epidermis of the hyperaccumulator species Hybanthus floribundus [44], T. montanum [60], Senecio coronatus [40], T. goesingense, Alyssum bertolonii, and A. lesbiacum [61], as well as other Alyssum species [64]. Severne [44] suggested that Ni accumulation in the epidermis of Hybanthus floribundus would decrease the cuticular transpiration; such change is considered as a xeromorphic adaptation. Nickel distribution in various epidermal cells was very far from uniform: in T. montanum leaves, Ni was found mostly in the auxiliary cells, whereas it was absent from stomatal guard cells and long basic cells [60]. According to other data, the oblong basic epidermal cells of T. goesingense leaves contained more Ni than the small cells surrounding the stomatal complex and the guard cells. The mechanisms underlying such distribution and its physiological meaning are unknown. As a whole, the hyperaccumulators T. goesingense, A. bertolonii, and A. lesbiacum accumulated Ni mostly in the vacuoles of the leaf and stem epidermal cell. The dimethylglyoxime method discerned small amounts of Ni at the base of branching stellate trichomes in Alyssum species [61, 64]. In opposite to these hyperaccumulators, Sebertia acuminata plants accumulated Ni mostly in the conducting tissues of their stems and in the fruits to the level of 1.15% per dry wt basis in the phloem and 0.13%, in the xylem. In the shoots, latex tubes in the phloem were stained most heavily with dimethylglyoxime [45]. Nickel accumulation in the conducting tissues was also reported in Quercus ilex. Here the pattern of Ni distribution was same as in Sebertia acuminata: the phloem accumulated more metal than the xylem [62]. In the fruits of S. acuminata, Ni accumulation decreased in the following order: the rudimentary endosperm > fleshy tissue > cotyledons of the embryo > fruit coats > seed coats [45]. Hence, many hyperaccumulators store Ni in covering and conducting tissues; as stated above, such localization would protect plants against the pathogens and herbivores and, as some authors believe [30, 42, 44], against drought. Nickel accumulation in covering tissues could also arise from plugged transpiration flow. Such suggestion is supported by the evidence that other RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL 263 Table 2. Ni distribution in plant organs Plant species Salt and its concentration in the medium Distribution in plant organs Source NiSO4 , 0.003–3 × 10–3 M shoots > roots [50] –3 NiSO4 , 0.003–3 × 10 M roots > shoots [50] NiCl2 ⋅ 6H2O, 25 mg Ni/kg soil roots > shoots (at tillering and booting stages); [52] generative organs > vegetative organs (at booting and milky and full maturity stages); the maximum in the caryopses at the full maturity stage Cajanus cajan NiSO4 ⋅ 6H2O, roots > shoots [54] 0.5 and 1.5 × 10–3 M Ceratophyllum demersum in water: 0.01–0.02 mg/kg, accumulates more metal than floating [55] in silt: 23–36 mg/kg Nymphoides flava Chenopodium ambrosioides NiSO4 , 50, 100 mg/kg soil roots > shoots [53] Cyperus difformis NiSO4 , 50, 100 mg/kg soil roots > shoots [53] Digitaria sanguinolis NiSO4 , 50, 100 mg/kg soil roots > shoots [53] Glycine max NiCl2 , 10–3–10–7 M nodules > roots > leaves > stems [17] Hybanthus floribundus 400 mg/kg soil leaves > photosynthesizing stems > lignified [44] stems > roots, flowers, and leaves > seeds Lolium perenne Ni(NO3)2 , 2–8 × 10–5 M shoots > roots [48] Myriophyllum spicatum in water: 0.01–0.02 mg/kg, accumulates more metal than floating [55] in silt: 23–36 mg/kg Nymphoides flava Nymphoides flava in water: 0.01–0.02 mg/kg, accumulated less metal than submerged Cerato- [55] in silt: 23–36 mg/kg phyllum demersum and Myriophyllum spicatum –3 Oryza sativa 0.5 × 10 M NiCl2 shoots > roots [56] Phaseolus vulgaris 2.2–3.75–7.5 × 10–3 mg/kg soil at high concentration: roots = pods > upper [26] leaves > lower leaves; at medium concentration: pods ≥ roots > leaves Psychotria douarrei not determined older leaves > young leaves [43] Sebertia acuminata 0.7% or 7 g/kg soil stems > leaves > fruits [45] Streptanthus poligaloides 2359–3840 mg/kg soil leaves > flowers > roots > seeds [57] Thlaspi arvense Ni(NO3)2 , 1–10 × 10–5 M roots > shoots [51] Thlaspi goesingense Ni(NO3)2 , 1–50 × 10–5 M shoots > roots [51] Triticum aestivum NiSO4 , 1 × 10–3 M roots > shoots [58] Typha latifolia Ni-EDTA, roots > shoots [29, 59] 10–150 and 600 mg/kg older leaves > young leaves Vigna unguiculata Ni-EDTA, 3.3 × 10–6 M older leaves > young leaves [10] Zea mays Ni(NO3)2 , 2–8 × 10–5 M roots > shoots [48] Plants of inaudated and meadow about 20 mg/kg soil roots > shoots [27] phytoceneses in Baikal region Alyssum lesbiacum Alyssum montanum Avena sativa heavy metals are non-specifically accumulated in the leaf epidermis and trichomes [36]. With the predominant Ni uptake via roots, it is of primary importance to outline the pattern of Ni distribution in the underground organs. A modified dimethylglyoxime method was used to demonstrate that Ni (35 µM) was present in all tissues of maize root following two days of exposure. Independent of root region and the particular tissue under study, Ni content in cell protoplasts exceeded that in the cell walls with the RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 highest metal concentration in the endoderm and pericycle. It follows that the endoderm does not block Ni translocation into the stelar tissues, as in the case of Cd and Pb. After 7-day exposure, Ni content considerably increased in all root tissues as compared to the data from 2-day exposure; however, the general pattern of Ni tissue distribution was not changed. In the longitudinal root sections, the authors observed large clusters of Ni-dimethylglyoximine crystals at the xylem vessel perforations. Although the cause of this phenomenon is No. 2 2006 264 SEREGIN, KOZHEVNIKOVA Table 3. The relative content of various nickel complexes in the hyperaccumulator plant species, % Species Casearia silvana Hybanthus floribundus Lasiochlamys peltata Phyllanthus serpentinus Psychotria douarrei Rinorea bengalensis Xylosma vincentii 2+ Ni ( H 2 O ) 6 33 5 33 18 21 7 36 Ni-citrate Ni-malate 67 95 67 42 16 93 64 – – – 40 63 – – Note: Data from [69]. (–) Not detected. not known, such evidence indicates the probable mechanism for limiting Ni transport into the shoots in the excluder plant species [49]. Nickel accumulation in the cell walls and the vacuoles in the cortical cells of onion roots was also established with the electron microscopical technique [65]. To conclude, Ni distribution in plant tissues is different from the pattern characteristic of such heavy metals as Cd and Pb. While the endoderm limits the movement of the latter into the central cylinder, Ni is freely translocated into the stelar tissues and in this way can easily reach the above ground organs of the accumulator plants, which might partly determine a specificity of its toxic effect. Ni TRANSPORT IN THE PLANT The pattern of Ni accumulation in cell walls and protoplasts of all root tissues presumes that Ni is translocated via both the apoplast and symplast. When reaching the xylem by radial transport, Ni will reach the shoots with the xylem sap [25, 45, 48, 51, 66, 67]. Within the late 25 years, numerous authors established that in addition to its ion form [45], Ni is translocated from roots to shoots in several forms [25, 45, 50, 66, 68–72], with citrate and malate complexes most widely distributed among them [45, 68, 70]. The chromatographic and spectrometric techniques demonstrated that the Ni–citrate complexes prevailed over the Ni–malate complexes in the leaves of such hyperaccumulator plants as Sebertia acuminata, Homalium francii, H. guillainii, H. kanaliense, Hybanthus austrocaledonicus, H. caledonicus, and Dichapetalum gelonioides [25, 68, 70], apparently due to higher stability of the former: log K [ Ni ( cit )– ] = 5.47 as compared to logK[Ni (mal)] = 3.30 [70]. Beside Ni complexes with organic acids, several 2+ plant species comprise aquacomplexes Ni ( H 2 O ) 6 [68, 69] and Ni complexes with histidine [50, 71, 72] based on high Ni2+ affinity for imidazole [1]. The con- tents of these complexes considerably vary in diverse plant species [69] (Table 3). The views on the role of histidine in binding Ni in the hyperaccumulator plants are rather inconsistent. When the hyperaccumulator plants Alyssum lesbiacum were grown on 0.3 mM Ni(NO3)2 solution, the histidine content in the xylem sap increased 36-fold; nothing of the kind occurred in the susceptible to Ni2+ A. montanum plants under the same conditions [50]. Similar increase in histidine content was reported in Brassica juncea [72]. It is of interest that spraying the A. montanum leaves with histidine promoted plant tolerance to nickel [50], and these authors hypothesized that histidine was involved in the mechanisms of A. lesbiacum tolerance to high nickel content in the shoots. The opposite pattern was observed when another hyperaccumulator, Thlaspi goesingense, was grown on 50 µM Ni(NO3)2 solution: the content of free histidine did not increase in the roots, shoots, and xylem sap as compared to the non-accumulating species, T. arvense. The content of Ni–histidine complexes in the shoots of T. arvense even exceeded by 5–10 times that in T. goesingense within the initial seven days of exposure to 10 µM Ni(NO3)2 solution. When Escherichia coli cells were transformed with the T. goesingense genes THG1, THB1, and THD1, which encode such enzymes for histidine biosynthesis as ATP-phosphoribosyl transferase, imidazole glycerolphosphate dehydrogenase, and histidinol dehydrogenase, the expression of these genes was not affected by Ni2+. In addition, the content of the corresponding mRNAs in T. goesingense roots and shoots did not depend on Ni2+ content in the nutrient solution. It follows that in T. goesingense, histidine does not contribute to the mechanisms of nickel hyperaccumulation and detoxification, although it may play some part in nickel translocation [71]. The amount and distribution of Ni-containing complexes would vary depending on plant age. When Glycine max plants were grown on the nutrient solution containing 0.05 µM 63NiCl, the highest content of Ni complexes with amino acids or peptides in exudate was registered at early growth stages, while the content of organic acids increased progressively with plant age [66]. The long-distance Ni transport may involve both the xylem and phloem. Thus, in Ricinus communis, a Nicontaining complex with polynucleotide or nucleoproteid as an organic component was found in 3–5 days following the introduction of 63Ni2+ to the nutrient solution (cited after [25]). The possibility of Ni translocation from the xylem to phloem was also demonstrated in shoots and glumes of intact girdled shoots of Triticum aestivum [67]. The rate of long-distance metal transport is different in various plant species. By lowering the rate of transporting metals, the excluder plants can alleviate their toxicity. Thus, the rate of Ni xylem transport from the roots into the shoots of Lolium perenne exceeded 2–7-fold that in Zea mays and progressed in RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL both plant species as the metal concentration increased in the nutrient solution. As a result, under sublethal concentrations of Ni salt (under 40 µM), maize was more tolerant to Ni than ryegrass. In addition, the analysis of xylem exudates of L. perenne shoots and Z. mays roots showed that the exudates were enriched with organic acids, such as malate, citrate, oxalate, and cis-aconitate [48]. Binding metal ions in the xylem sap may turn out a nonspecific mechanism of detoxifying and transporting heavy metals providing for accumulation and translocation without evident manifestations of toxicity. In its turn, the efficacy of the chelating system depends on plant species and its specific metabolism as well as on the affinity of metal ions for particular chelating agents. The latter characteristic is determined by the stability constant of the corresponding complex. For many years, the hyperaccumulators were believed to manifest high rates of metal translocation. However, similar rates of metal transportation from roots to shoots were observed in tolerant T. goesingense and susceptible T. arvense plants grown for 12 h on the nutrient solution with nontoxic Ni2+ concentration of 10 µM [51]. It follows that the capacity of the hyperaccumulator species to concentrate Ni in their shoots relies on more efficient detoxification mechanisms than in non-hyperaccumulators. Below we will consider the intracellular aspects of such detoxification. INTRACELLULAR NICKEL LOCALIZATION In the hyperaccumulator species Alyssum serpyllifolium, A. bertolonii, A. lesbiacum, and Thlaspi goesingense, Ni is predominantly localized in the vacuoles as the complexes with organic acids; as a result, most of Ni is removed into a metabolically inactive compartment; such compartmentation exemplifies a nonspecific mechanism of heavy metal detoxification [36, 51, 61, 73, 74]. When A. serpyllifolium plants were grown from seeds for 6 weeks in the presence of 500 µg Ni(NO3)2/g soil, the highest Ni content was registered in the vacuoles as malate and citrate complexes. Considerably lower amount of complexes was found in the cell walls (3.6%), chloroplasts (1.0%), mitochondria (2.1%), and microsomes (3.0%) [73]. Meanwhile in the case of other hyperaccumulator, Hybanthus floribundus, high Ni content was found in the cell walls wherein Ni was bound to pectins [40]. Binding metal ions to the cell wall components depends on the affinity of particular metal ions for polygalacturonic acid [75, 76]. The capacity of cell walls for binding metals may shape plant tolerance towards heavy metals. The capacity to accumulate Ni in the vacuoles as the complexes with organic acids also determines plant tolerance towards high levels of Ni in the environment. Thus, the protoplasts isolated from T. goesingense leaves were more tolerant to nickel sulfate (250 µM) than those from the susceptible species T. arvense [51]. RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 265 Using the X-ray spectroscopy, these authors demonstrated that, following 24-h exposure of the seedlings of these two species to 1 µM NiSO4 solution (labeled with 40 µCi 63NiCl2), Ni content was similar in the apoplast (65–70% of the total content), whereas Ni content in the vacuoles of T. goesingense (15–20%) exceeded the corresponding index in T. arvense (about 8%). Apparently, the Ni compartmentation in the vacuoles made the former plant species more tolerant to it [74]. These data are in contrast with the later evidence from the experiment with the hyperaccumulators T. goesingense, A. bertolonii, and A. lesbiacum. Thlaspi plants were grown in soil culture with addition of 2 g Ni (as NiSO4) per kg soil for 86 days, Alyssum plants were grown for 64 days with Ni content of 4 g/kg soil [61]. The X-ray microanalysis of these plants established the predominant Ni accumulation in the vacuoles of leaf epidermal cells. Ni contents were lower in the cell wall and cytoplasm. The species-specific variations in the intracellular Ni distribution may arise from dissimilar salt concentrations and exposure periods. Indeed, Kramer et al. [50, 51] analyzed the leaves containing less than 1 g Ni per kg dry wt, whereas Kupper et al. [61] used the leaves with a tenfold higher Ni concentration; in the latter case, with the Ni content typical of hyperaccumulator plants, the experimental data seem better validated. In addition, the assessment technique was more sensitive in the latter case. The transfer of metal ions or their complexes into the vacuole may rely on the characteristic protein carriers located in the tonoplast. Such carriers were found in T. goesingense and named TgMTPs (Thlaspi goesingense Metal Tolerance Proteins). Proteins TgMTP1t1 and TgMTP1t2 are coded by the common DNA sequence TgMTP1 and differ in the amino acid sequence of the histidine-rich metal-binding domain, which affects the metal specificity of these carriers. Thus, TgMTP1t1 expression in transformed yeast cells provided for the tolerance towards Cd, Co, and Zn, whereas the expression TgMTP1t2, towards Ni. As compared to non-accumulating species T. arvense, Arabidopsis thaliana, and Brassica juncea, the hyperaccumulator T. goesingense maintained high level of TgMTP1 expression promoting Ni transfer into the vacuole and providing for the successful Ni detoxification [77]. There are conflicting data as to the role of metal/H+ antiport in heavy metal transfer across the tonoplast [78– 80]. The study of Ni transport into the vacuoles of Avena sativa roots demonstrated that the tonoplast lacked both Ni2+/H+-antiport and N-nucleotide-dependent pump [79]. In the hyperaccumulators A. murale [79] and A. lesbiacum [81], a Ni2+/H+-antiport on the tonoplast provided for rapid Ni2+ uptake into the vacuole. By the predominant accumulation in cell protoplasts, Ni fundamentally differs from Cd and Pb, which are mainly bound by cell wall components. Hence, the problem of Ni transport across the plasmalemma into the cell demands further research. The formation in the vacuoles of the low-soluble Ni complexes with organic No. 2 2006 266 SEREGIN, KOZHEVNIKOVA Table 4. Effect of nickel on enzyme activity Concentration Ni, mM Enzyme activity CO2 fixation Calvin cycle 0.5; 1 0.5; 1 ↓ ↓ Cajanus cajan Cajanus cajan [84] [84] Calvin cycle Calvin cycle Calvin cycle Calvin cycle 0.5; 1 0.5; 1 0.5; 1 0.5; 1 ↓ ↓ ↓ ↓ Cajanus cajan Cajanus cajan Cajanus cajan Cajanus cajan [84] [84] [84] [84] ↓ ↑ ↓ ↓ ÇÂta vulgaris Oryza sativa ÇÂta vulgaris Glycine max [85] [86] [85] [87] <0.05 >0.05 0.01–1 ↑ ↓ ↑ Oryza sativa [21] [88] <0.05 >0.05 ↑ ↓ Allysum maritimum A. argenteum Oryza sativa [21] 0.01 ↓ Allysum maritimum [88] 0.1 <0.05 >0.05 1–40 ↑ ↑ ↓ ↑ Oryza sativa [21] Triticum aestivum [89] Enzyme Rubisco Glyceraldehyde 3-phosphate dehydrogenase 3-Phosphoglycerate kinase Aldolase Fructose 1,6-bisphosphatase NADP- and NAD-dependent phosphoglyceraldehide dehydrogenases Nitrate reductase H+-ATPase Glutamine synthetase Alanine aminotransferase Process IAA oxidase nitrate reduction ion transport glutamine synthesis transformation of alanine into pyruvate IAA oxidation Glutathione reductase glutathione reduction Ascorbate exidase ascorbate oxidation Superoxide dismutase O 2 deactivation Catalase H2O2 degradation Peroxidase polyphenolic oxidation .– 1 0.5 1 0.2 Plant species Source Note: ↓ and ↑—decrease and increase of enzyme activity. acids can be seen as a detoxification mechanism; besides, the tolerance of hyperaccumulator species may depend on the efficiency of the tonoplast carriers that account for Ni entry into the vacuole. EFFECTS OF NICKEL ON VARIOUS ENZYME ACTIVITIES Same as other heavy metals, Ni affects various physiological processes in plants, starting from several enzyme activities [36, 82, 83] (Table 4). It is not easy to discern direct and indirect effects of metals on enzyme activities. The latter effects arise from ion imbalance due to the competitive inhibition of absorption and transport of such nutrients as Zn, Fe, Cu, and others. Heavy metals may directly inhibit enzymes by interacting with protein SH-groups; in this way protein conformation is changed, and enzymes are inactivated. Presently, about a hundred of enzymes are known that are inhibited by SH-group binding, with concomitant metabolic disorders. When Zea mays seedlings were incubated on the solutions of Ag, Cd, Pb, Zn, Cu, Tl, Co, and Hg salts (0.001–3 g/l), the affinity of metals for SH-groups was found to significantly correlate with the molar concentrations inhibiting growth by 50% [90]. Apparently, Ni2+ binding to SHgroups is one of the mechanisms of in vitro Ni toxicity towards Mg2+-dependent ATPases of the plasmalemma; however, Ni2+ may directly bind to ATP and in this way deplete the substrate pool of ATPase [91]. The affinity of Ni2+ for sulfhydryl groups is lower than of other heavy metals. On the contrary, Ni affinity for histidine exceeds that of Cd and Pb, and therefore the inhibition of enzyme activity by nickel may result from the interaction with this ligand. The toxic effects of metals on enzyme activity in vitro do not always agree with the in vivo effects at the same salt concentration. Such disagreement may stem from the presence of efficient cellular mechanisms for detoxification and the physiological barriers that curb metal translocation into the cytoplasm. To illustrate, Ni2+ was RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL 267 Table 5. Effect of Ni on the contents of macro- and micronutrients in plants Plant species Plant organ Hordeum vulgare Oryza sativa Phaseolus vulgaris Lolium perenne Triticum aestivum Triticum aestivum (thillering phase) Triticum durum (thillering phase) Thlaspi montanum shoots shoots and roots roots leaves shoots leaves and roots leaves roots leaves roots leaves Salt/metal concentration 6 × 10–7 and 1 × 10–6 M NiSO4 5 × 10–4 M NiCl2 3.4–14 × 10–6 M Ni(NO3)2 ⋅ 6H2O 30, 90, 180, 270 mg Ni/kg soil 5–40 × 10–3 M NiSO4 ⋅ 7H2O 6.7 × 10–5 M Ni 6.7 × 10–5 M Ni soil (at the ambient conditions) N P K Ca Mg Fe Mn Cu Zn Source – – – – – ↓ ↑↑ ↑↑ ↑ – – 0 – – – – – ↓ – – – – – – – – – – – – – – ↑ – ↓ 0 0 – – ↑ 0 0 0 ↑ – ↓ – – – ↓ – – – – 0 ↑ 0 ↑↑ 0 0 0 0 – 0 – ↑↓ ↓ – ↓ – – – – – ↑ ↓ ↓ 0 ↓ 0 0 ↓ 0 0 0 ↓ ↓ 0 0 0 0 ↓ 0 0 – – – – – [4] [56] [92] [93] [89] [94] [94] [60] Note: 0—No changes in uptake; ↑—increased uptake; ↓—decreased uptake; (–) no data available. shown to promote in vivo Mg2+-dependent ATPases in the plasmalemma of Oryza sativa shoots [86]. Total decline of enzyme activities is sometimes observed due to decreased enzyme contents. Thus, the decrease in nitrate reductase activity in soil-grown Beta vulgaris plants following the addition of 1 mM NiSO4 resulted from the diminished rates of nitrate uptake and translocation into the shoots wherein nitrate is reduced. Nitrate in the cytoplasm induces the expression of the nitrate reductase gene, and hence it is the shortage of nitrate in the cells that would primarily decrease the enzyme concentration. Besides, glutamine synthetase and alanine aminotransferase activities were also lowered in this case; both activities considerably depend on the cytoplasmic levels of nitrate and their substrates [85, 87]. Similar mechanism of indirect influence on nitrate reductase activity was established for other heavy metals [36]. Depending on its concentration, nickel ion can both stimulate and inhibit enzyme activities in plant tissues (Table 4). Thus, the activities of IAA oxidase, ascorbate oxidase, catalase, and peroxidase in O. sativa seedlings were at their highest at 50 µM NiCl2; the enzyme activities considerably declined at a higher Ni concentration and were promoted at a lower Ni concentrations. In addition, it was shown that Ni did not affect these enzymes directly: under in vitro conditions, the same Ni concentrations did not produce any visible inhibition of enzyme activities [21]. Enzyme in vivo resistance to Ni varies as related to plant development stage. Thus, in Cajanus cajan leaves collected from young plants (30 days after sowing), Ni inhibited the activities of the Calvin cycle enzymes (Table 4). Meanwhile, the inhibitory effect was inconsiderable when Ni salt was added at the later developmental stage (70 days after sowing) [84]. However, the mechanism of this phenomenon is unknown. The production of reactive oxygen species in plant cells is another universal mechanism of heavy metal RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 toxicity. Plants respond to oxidative stress by elevating the activity of the antioxidant enzymes of the ascorbate–glutathione cycle, such as catalase, peroxidase, superoxide dismutase, glutathione reductase, and ascorbate oxidase, which protect plant cells against free radicals [21, 36, 88, 89]. The effect of Ni2+ on antioxidant enzyme activities may differ in the accumulator and non-accumulator plant species. When non-accumulator A. maritimum plants were grown on the nutrient solution with Ni added, the activities of superoxide dismutase, ascorbate peroxidase, and glutathione reductase were enhanced, whereas these activities were diminished in the hyperaccumulator A. argenteum, while that of superoxide dismutase was inhibited [88] (Table 4). The tolerance to Ni in the hyperaccumulators seems to employ other mechanisms of effective Ni2+ detoxification; in this way, Ni content is diminished in the cytoplasm, and the demand for antioxidant enzyme activities is alleviated. All these data led us to conclude that, at high Ni concentrations, most of enzyme activities were diminished, whereas some activities, especially those of the antioxidant enzymes, increased. In most cases, we do not know whether these changes in enzyme activities stem directly from Ni2+ effects, such as binding to SH-groups or histidine or displacing the metals from metal–enzyme active centers, or indirectly, when mediated by the chain of reactions that affect the expression of the corresponding genes or exhaust their substrate pools. The inhibition of enzyme activities by heavy metals is one of the causes of declining cell metabolism. EFFECT OF NICKEL ON MINERAL NUTRITION Heavy metals curb cation and anion absorption by plant roots; this is one of nonspecific mechanisms of their toxicity [36]. However, the published evidence on the effects of Ni on plant mineral nutrition is rather contradictory (Table 5). No. 2 2006 268 SEREGIN, KOZHEVNIKOVA In the presence of Ni, the contents of mineral nutrients in plant organs may increase, decrease, or stay even (Table 5). One of the probable mechanisms for decreasing the uptake of macro- and micronutrients relies on the competition for the common binding sites due to the comparable ionic radii of Ni2+ and other cations. Such mechanism may operate [94] when the uptake of Mg2+ (78 pm), Fe2+ (82 pm), and Zn2+ (83 pm) is decreased in the presence of Ni2+ (78 pm) (ionic radii in parentheses are from [23]). One should emphasize that the lowered uptake of Mg and Fe is one of the causes of chlorosis produced by the excess of Ni in the environment [92, 93]. The decline in nutrient uptake may also result from the Ni-induced metabolic disorders that affect the structure and enzyme activities of cell membranes [36]. Thus, Ni2+ affected the sterol and phospholipid composition of the plasma membrane in Oryza sativa shoots, with concomitant changes in the ATPase activity [86]. Apparently, these changes affected the membrane permeability and in this way changed the ion balance in the cytoplasm. The effects of Ni on nutrient uptake depend in many aspects on Ni concentration in the environment. The experiments with ryegrass plants demonstrated that Fe content in the shoots increased at low Ni concentrations and decreased at higher concentrations [93] (Table 5). An increase in soil Ni content from 50 to 200 mg/kg soil decreased the contents of Cu and Mg in the caryopses and Mg and Ca in the shoots of Triticum aestivum [94]. The data presented in Table 5 allow us to conclude that at high Ni concentrations (about 0.1 to 1 mM), the contents of macro- and micronutrients in plant tissues are usually lowered down because of disordered absorption and transport [56, 89]. At the same time, at low Ni concentrations in the environment (10 to 1 µM), the contents of nutrients did not change and in some cases even increased [92, 94]. Such phenomenon was described as the concentrating effect; these effects are seen as the result of growth inhibition (dry biomass decreases) in the plants grown on the nutrient solutions low in Ni, while the rate of metal absorption stays the same as in the control plants; consequently, the contents of heavy metals increase per unit of dry matter [92]. The effects of the same Ni concentration would vary in diverse plant species. Thus, when the plants of T. aestivum and T. durum were kept on 67 µM Ni, the contents of Ca and Mg increased and that of Zn decreased in the leaves of the former species and did not change in the latter [94] (Table 5). The toxic Ni concentrations specifically affect the ionic balance in various plant organs. The contents of Fe, Mn, and Zn decreased in T. aestivum leaves at the tillering stage, while only Mn content declined in the roots [94] (Table 5). Plant species tolerant and susceptible to Ni may differ by the changes in their mineral contents as affected by Ni. When two cultivars of T. aestivum were grown in the soil contaminated with Ni (50–200 mg/kg soil), Fe concentration in grain declined in both cultivars and the contents of Cu, Ca, and Mg, only in the susceptible genotype. The latter also manifested Mn and Mg deficiency in the leaves, apparently resulting in chlorosis [94]. This evidence and some additional data presume that the resistance of transport systems to heavy metals provides one of the mechanisms of plant tolerance. EFFECT OF NICKEL ON WATER REGIME The stability of plant water regime depends on the balance between water uptake and transpiration. Many authors reported that Ni induced the decline in plant transpiration and water content [84, 85, 88, 95, 96]. Following four days of growth of Triticum aestivum plants in the sand culture, with 10 mM Ni added to the nutrient solution, leaf water potential, stomatal conductance, the transpiration rate, and total moisture content decreased, especially in the uppermost leaf wherein the metal accumulation was most pronounced [95]. Transpiration may decline as a consequence of several metal-induced changes that are also produced by other heavy metals. First, the toxic effect of Ni2+ on plant growth would decrease the area of leaf blades, the major transpiring surface. Such decrease of leaf area by 40% was observed in Cajanus cajan plants grown in sand with 1 mM NiCl2 added to the nutrient solution [84]. Similarly the leaf area was diminished in Brassica oleracea plants grown in agar in the presence of 5–20 g/m3 NiSO4 ⋅ 7H2O [96]. Second, transpiration may decrease because of lower stomata numbers per unit of leaf area [96]; nonetheless in some cases, stomata density may even increase due to the reduction of leaf area and the size of epidermal cells [97]. The induction of stomata closure is among the primary effects of heavy metals [36, 96]; such closure would also diminish transpiration. In addition, damaged and therefore permanently closed stomata were found in B. oleracea [96]. The presence of Ni in Phaseolus vulgaris leaf tissues was shown to elevate the level of ABA, which is known to induce stomata closure (cited after [95]). The decrease in moisture content and stomatal conductance induced by Ni is also one of the mechanisms of its toxicity towards photosynthesis; we will deal with this phenomenon below. EFFECT OF NICKEL ON PHOTOSYNTHESIS For heavy metals, several direct and indirect ways are known to lead to nonspecific inhibition of photosynthesis. The diminished rate of photosynthesis is related to disrupted chloroplast structure, blocked chlorophyll synthesis, disordered electron transport, inhibited activities of the Calvin cycle enzymes, and CO2 deficit caused by stomatal closure [36]. RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL ADP 269 + Pi Cd, Ni ATP Cd, Ni, Pb Cd NaDPH Cd, Ni, Pb Cd hν Ni NaDP + β hν Stroma 2 H+ D2 D1 e– PQ FNR C E I A PQH2 P680 e– MSP RFeS Cytb6 f e– PC γ ε n H+ I J e– N II CF2 P700 G β B PSI PSII α Fdx– Fdx D σ CF1 2 H+ III F PC– Cd, Cu H2O 1/2O2 + 2 H + 2 H+ Ni, Cu Pb Thylakoid lumen Fig. 1. Effect of heavy metals on the light-dependent reactions of photosynthesis. MSP—the photosynthetic system of water photolysis involving manganese stabilizing protein; P680—the reaction center of PSII; D1 and D2—PSII proteins; PQ and PQH2—oxidized and reduced plastoquinone pools; RFeS—Fe-containing Rieske-type proteins; PC—plastocyanin; P700—the reaction center of PSI; A to N—PSI proteins; Fdx—ferredoxin; FNR—ferredoxin–NADP+ oxidoreductase; CF0–CF1—ATPase particles. Arrows show the pathways of electron and proton transport. The data for Cd, Pb, and Cu are compiled from [36, 100, 101]. The Z scheme of the membrane organization follows [102]. The decrease in chloroplast size and numbers and the disorganization of chloroplast ultrastructure, including the diminished numbers of grana and thylakoids, their deformation, the formation of plastoglobuli, and the changes in the membrane lipid composition, were reported in Brassica oleracea plants grown in agar in the presence of NiSO4 ⋅ 7H2O (10–20 g/m3). Such changes seemed to arise from the Ni-induced decline in cell moisture content or from an oxidative stress resulting in peroxidation of membrane lipids [96]. Several authors reported diminished chlorophyll content in the leaves of Ni-treated plants; such chlorosis could result from both Fe and Mg deficiency and the inhibition of chlorophyll synthesis [53, 84, 92, 93, 96]. The disruption of electron transport exemplifies another mechanism that brings down the photosynthetic production. Numerous experiments demonstrated that Ni2+, similar to other heavy metals, primarily affects PSII [36, 98–101]; this evidence is in line with the predominant Ni accumulation in the PSII-containing lamella regions [98]. When inspected in more detail, Ni was shown to inhibit electron transport from pheophytin via plastoquinone QA and Fe to plastoquinone QB by changing the structure of carriers, such as plastoquinone QB, or the reaction center proteins [99, 100]. In the thylakoids, RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 Ni ions also decreased the contents of cytochromes b6f and b559, as well as ferredoxin and plastocyanin; as a result, the efficiency of electron transport dropped down [98]. Figure 1 illustrates schematically a generalized pattern of heavy metal effects on the light photosynthetic reactions. By inhibiting key enzyme activities of the Calvin cycle, such as Rubisco, 3-phosphoglycerate kinase, fructose-1,6-bisphosphatase, aldolase, and NAD- and NADP-dependent phosphoglyceraldehide dehydrogenases, heavy metals can also hold back the dark reactions of photosynthesis. Such effects were demonstrated in Cajanus cajan leaves following several days of incubation on 1 mM NiCl2 solution [84]. The blockade of the Calvin cycle reactions would lead to the accumulation of ATP and NADPH produced by the light reactions; the latter, in their turn, develop a high pH gradient across the thylakoid membrane that blocks the PS II activity [100]. Another mechanism of curbing the photosynthetic productivity stems from the stomatal closure in Ni-stressed plants that limits plant CO2 uptake [84, 95]. The toxic effects of heavy metals on many other metabolic processes would amplify the direct inhibition of photosynthesis. All these metabolic changes inhibit plant growth and disrupt morphogenesis; the ensuing No. 2 2006 270 SEREGIN, KOZHEVNIKOVA Table 6. The range of metal toxicity in several plant species Plant species Hordeum vulgare Lolium perenne Triticum aestivum Vicia faba Zea mays Range of metal toxicity Source Hg > Pb > Cu > Cd > Cr > Ni > Zn Cu > Ni > Mn > Pb > Cd > Zn > Al > Hg > Cr > Fe Cu > Cr > Ni > Zn > Pb ≈ Cd > Al > Fe Cd > Ni > Zn ≈ Co Cu ≈ Tl > Ag > Cd > Hg > Co > Zn > Pb or Tl3+ > Cu2+ > Ag+ > Hg2+ ≈ Cd2+ > Ni2+ > Zn2+ ≈ Pb2+ ≈ Co2+ > Sr2+ [107] [104] [108] [108] [90] Table 7. Nonspecific mechanisms of toxicity, detoxification, and hyperaccumulation of heavy metals Toxicity Detoxification 1. Inhibition of enzyme activity 1. Binding metals to cell wall components 2. Development of reactive oxygen species and oxidative 2. Transport and accumulation stress of metals in vacuoles as low solubility complexes with organic acids 3. Inhibition of cation and anion uptake 3. Metal binding by chelators in xylem sap 4. Changes in membrane permeability due to the changes 4. Decreased metal transport from in membrane lipid composition and the inhibition of mem- roots into shoots brane enzyme activities 5. Changes in water regime due to declining water poten- 5. Enhanced activity of antioxitial, stomatal conductance, the transpiration rate, and total dant enzymes water content 6. Decline in photosynthesis rates due to obstructed elec- 6. Synthesis of osmolytes, such as proline and polyamines tron transport, inhibited synthesis of chlorophyll and Calvin cycle enzymes, and CO2 deficiency brought about by ABA-induced stomata closure 7. Decline in respiration rate 7. Metal exclusion into the root slime 8. Inhibition of growth and morphogenesis due to decreased plasticity of cell walls that resulted from cell wall lignification, hindered mitosis, and chromosomal aberrations 9. Morphological and anatomical changes at various levels of organization, such as decrease in the intercellular spaces; mesophyll cell volumes; the sizes of conducting bundles, the changes in chloroplast structure, etc. phenomena are often used to assess the phytotoxicity of heavy metals. EFFECT OF NICKEL ON PLANT GROWTH AND MORPHOGENESIS The toxic effects of nickel and other heavy metals are primarily manifested as the inhibition of plant growth [36, 49, 53, 54, 56, 58, 59, 86, 89, 93, 103, 104], an index widely employed to assess the environmental pollution [105]. Growth inhibition gains strength at higher metal concentration in the medium [105, 106]. In the excluder species, which accumulate Ni mostly in their roots, root growth is inhibited more Hyperaccumulation 1. Efficiency of ion absorption and uptake 2. Resistance of plant transport systems to heavy metals 3. Enhanced metal-binding capacity of cell walls 4. Efficiency of the mechanisms for metal detoxification and maintaining homeostasis heavily than the growth of shoots [49, 54, 104, 106], and therefore the root test is widely used for evaluating the toxicity of various agents, including heavy metals [36, 49, 104, 105]. The tolerance index, that is the ratio (%) between the root/shoot length of the heavy metalstressed plant and that of the control plant [54, 106], and LC50, the metal concentration that inhibits root growth by 50%, are the indices of plant tolerance toward heavy metals [104, 105]. Using the latter index, Wong and Bradshaw, arranged the metals by decrease in their toxicity to root growth of Lolium perenne seedlings in the following order: Cu > Ni > Mn > Pb > Cd > Zn > Al > Hg > Cr > Fe [104]. This range will change in other plant species [107, 108], apparently because RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL CTR1 Cd, Zn, Ni Zn, Mn, Cd Cd Cu CAT POD SOD + PMP Vacuole Zn, Cn, Ni, Co TgMTP LCT1 Cd, Pb, Ni åÂ2+ Oxidative stress ROS MT ATP Cu, Zn, Cd ? + CRS5, CUP1 etc. Nucleus gsh1 + + Cd gsh2 Organic acids (malate, citrate, oxalate) Zn, Pb, Ni, Cd Cd, Ni ADP Cd Gene expression ? Zn, Cd, Ni, Co Me/H+antiport ZIP C‡-channel 271 + PC Cd Cu Zu Bi Hg Ag CAD1, AtPSC, TaPSC etc. Complex PC-å Cd HMT1* ? + ATP ATP-arylsulfurylase SO42– ATP APS PPi GCS Cys Glu ATP GS ADP + Pi PCS GSH γ-Glu-Cys ADP + Pi Gly ATP åÂ2+ ADP + Pi PC Complex PC-å – CYTOPLASM CELL WALL Binding to cell wall pectins Cd, Pb, Zn, Ni Fig. 2. The entry of heavy metals in the cytoplasm and the putative pathways of their detoxification. ROS—reactive oxygen species; APS—adenosine 5'-phosphosulfate; CAT—catalase; Cys—cysteine; Glu—glutamate; GC— γ-glutamylcysteine; GCS—γ-glutamylcysteine synthetase; GS—glutathione synthetase; GSH—glutathione; Gly—glycine; Me2+—bivalent metals; MT—metallothionein; PC—phytochelatine; PCS—phytochelatine synthase; PMP—phytometallophores (phytosiderophores); POD—peroxidase; SOD—superoxide dismutase. ZIP, LCT1, and CTR1 are metal transporters at the plasma membrane; TgMTP and HMT are metal transporters at the tonoplast. Putative processes are indicated with the question sign. (+) and (–) mean activation or inhibition. The processes established in Schizosaccharomyces pombe are marked with an asterisk. Data were compiled from [32–34, 36, 82, 83, 119–124, 126, 127]. plant species differ in their tolerance. However, one must emphasize that such ranges do not always reflect genuine toxicity as some authors used the weight rather than molar concentrations of metals [90] (Table 6). By comparing LC50 indices in diverse plant species, we classify them into more tolerant (Cucumis sativus and Panicum miliaceum) and less tolerant (Chloris gayana, Lactuca sativa, Lolium perenne, Panicum maximum, and Zea mays), with LC50 lower by the order of magnitude [104, 105]. Unlike root growth, the process of lateral root initiation is very resistant to most heavy metals [36, 109], due to the endodermal barrier and the characteristic structure of the central cylinder cells [110, 111]. However, Ni2+ considerably decreased the number of lateral roots in rice and maize [49, 106], apparently because Ni can cross the endodermal barrier and accumulate in the pericycle cells. Beside the root growth, Ni2+ reportedly exerts considerable inhibitory effect on shoot growth and morphogenesis in Phaseolus vulgaris [92], Digitaria sanRUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 guinolis, Cyperus difformis, and Chenopodium ambrosioides [53]. The total growth inhibition in P. vulgaris also affects seed formation [92]. Seed germination is the most resistant to heavy metals. The caryopses of rice [21] and maize [112] germinated at high concentrations of Ni salts, such as 10–2 M. In addition to toxic effects on growth, heavy metals may change plant morphology and anatomy. Thus, the exposition on 1 mM NiSO4 solution decreased the mesophyll thickness, the size of vascular bundles, the vessel diameter in the main and lateral vascular bundles, and the width of epidermal cells in Triticum aestivum leaves [58], whereas in the leaves of Brassica oleracea plants grown in agar in the presence of NiSO4 ⋅ 7H2O (10–20 g/m3), the volumes of intercellular spaces and palisade and sponge mesophyll decreased as compared to the control plants [96]. The mechanisms of inhibition of plant growth and development by Ni2+ are insufficiently clarified. In addition to general metabolic disorder, heavy metals are known to decrease the plasticity of cell walls, probNo. 2 2006 272 SEREGIN, KOZHEVNIKOVA ably by direct binding to pectins and by promoting peroxidase activity in the cell walls and intercellular space; these peroxidases are essential for lignification and linkage between extensin and polysaccharides containing ferulic acid [89]. Another way to inhibit growth is by hampering cell divisions [113–117]. The incubation on 1.5–5 mM NiCl2 solution brought down the mitotic index in Vicia faba roots [115], and at the concentration of 60 mM, in Zea mays roots [114]. At the concentration of 0.1 mM, NiSO4 blocked cell divisions in the rhizoderm, exoderm, middle cortex, except in the distal cells of these tissues, and in the peripheral cells of caliptrogen in the embryonic root of Triticum aestivum [117]. The inhibition of cell divisions was frequently accompanied by disorganization of nuclear structures. Thus, in the root tips of Cajanus cajan plants grown in the presence of 1.5 mM NiSO4 ⋅ 6H2O, two nucleoli developed in the nucleus, the chromatin became exceedingly condensed, and the nuclear membrane was disrupted [54]. Heavy metals may cause mitosis disorder and chromosome aberrations. In the meristematic cells of Allium cepa root, Ni2+ (10 to 100 µM) produced various chromosome aberrations: C-metaphases, sticky chromosomes, and chromosome bridges, while the interphase cells contained micronuclei. Besides, at high Ni2+ concentrations (1–10 mM), the nuclear material was found in the cytoplasm, whereas the nuclei contained nucleoli of irregular form (oval, oblong, and dumbbelllike). Similar changes were observed in plant cells exposed to other heavy metals; however, the damage extent depending on concentration became the basis for ranging the metals by decreased mutagenic effect (the minimum toxic concentration is listed in parentheses): Hg2+ and Cd2+ (10–7–10–5 M) > Zn2+, Pb2+, Cu2+, Ni2+, Co2+, Al3+ and Cr3+ (10–4–10–3 M) > Mn2+ and Mg2+ (10–2 M) [118]. To conclude, plant growth inhibition by nickel and other heavy metals results from general metabolic disorder and immediate inhibition of cell divisions. However, it is not clear whether Ni enters cell nuclei at high concentrations and, if it does, how important is immediate Ni interaction with DNA and nuclear proteins. The possible effect of Ni2+ on fragmoplast formation is also unknown. By elucidating these issues, we will better understand the toxic effects of nickel on plant growth and morphogenesis. CONCLUSION Our analysis of the published data on Ni2+ distribution, translocation, and toxic effects and on plant responses to nickel excess showed the characteristics that are peculiar for nickel or common with other heavy metals. The specific mechanisms of Ni2+ absorption by plant root systems have not been yet elucidated. Soil is the major source of nickel for plants, and similar to other heavy metals, Ni availability is higher in overmoistened soils of low humus content, light granulometric composition, and low pH of the soil solution. The ions of other metals decrease Ni2+ availability, although the effects of Ca2+ may vary. The great number of plant species that hyperaccumulate more than 1 g Ni per kg of dry shoots is a characteristic aspect of Ni distribution in plant organs. The researchers still debate over the causes and mechanisms of hyperaccumulation, which may depend on the morphological and physiological characteristics of particular plant species, the capacity of Ni to overcome the physiological barriers, etc. (Table 7). Plants comprise several such barriers that curb the entry of heavy metals into the shoots, primarily the plasmalemma and endodermis at the cell and tissue levels, respectively. However, these barriers are not universal: most of Cd and Pb are translocated via the apoplast, and their entry into the central cylinder is limited at nonlethal concentrations, while Ni2+ freely enters the stele via the symplast. The specific carriers providing for Ni uptake in plants have not been yet identified. The uptake of heavy metals into plant cytoplasm is mediated by various transport systems localized at the plasmalemma; some of them are shown in Fig. 2. In grasses, the uptake of Ni and other heavy metals was shown to employ Ca-channels and the phytometallophore mechanism. The intracellular localization of heavy metals and their distribution in plant tissues determine the profile of their toxicity affecting various physiological processes. Nickel exerts both general and specific toxicity. The former manifestations characteristic of most heavy metals include the disorders in mineral nutrition, water regime, photosynthesis, growth and developments, etc. (Table 7). The specific pattern of Ni toxicity is illustrated by the inhibition of lateral root development: it is in this aspect where Ni toxicity differs from that of other heavy metals, such as Ag, Cd, Pb, Zn, Cu, Tl, Co, and Hg, which blocked root growth at nonlethal concentration without inhibiting root branching. The inhibition of root branching by Ni stems from its accumulation in the endoderm and pericycle and the interruption of cell divisions in the latter. When heavy metals enter plant cells, they incite several specific and nonspecific systems of protection and detoxification, such as the immobilization in the cell walls and vacuoles or the induction of catalase, peroxidase, and superoxide dismutase, which account for the neutralization of free radicals and peroxides that are progressively accumulated in the course of metalinduced oxidative stress; other protection systems include the synthesis of osmolites, such as proline, the changes in the cell wall composition such as callose and suberin deposition, the imbalance of plant hormones, primarily ethylene and ABA, the synthesis of metallothioneins, phytochelatins, etc. (Table 7, Fig. 2). All these changes are the links in one and the same chain of events representing cell responses to the entry of heavy RUSSIAN JOURNAL OF PLANT PHYSIOLOGY Vol. 53 No. 2 2006 PHYSIOLOGICAL ROLE OF NICKEL metals and aimed at maintaining cell homeostasis. The accumulation of heavy metals in plant vacuoles as the complexes with organic acids is a universal mechanism of detoxification. The transport of heavy metals into vacuoles may employ various mechanisms, such as metal/H+-antiports, although this mechanism has not been recognized universally, and the tonoplast carriers, which transfer metal ions (TgMTP) or metal complexes with phytochelatins (HMT1) (see Fig. 2). The latter are small peptides synthesized in the cytoplasm from glutathione with the aid of the enzyme phytochelatin synthase. Cadmium ion is the strongest activator of this enzyme; Cd2+ also activates the expression of gsh1 and gsh2, which control the separate steps in glutathione biosynthesis [119–124]. As different from other heavy metals (Cd, Ag, Cu, Zn, Bi, and Hg), Ni does not activate the synthesis of phytochelatin synthase; as a result, phytochelatins are not of great importance for nickel detoxification [125]. Another mechanism of detoxification is employed to neutralize Cu2+ and Zn2+ ions: they induce the synthesis of metallothioneins, which, in contrast to phytochelatins, are the primary gene products. The entry of Cu2+ and Zn2+ ions into plant cells activates the expression of the corresponding genes CRS5, CUP1, etc. [124, 126, 127] (Fig. 2). In conclusion, we would emphasize that both toxic effects of heavy metals and their detoxification are the complex processes involving numerous related and interacting mechanisms. The efficiency of these mechanisms is probably the primary factor of plant tolerance and plant capacity for hyperaccumulation. 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