ARTICLE   Open Access    

Insights into physio-biochemical responses of maize to salt shock stress and removal of the stress at the whole-plant level

More Information
  • Timely increased antioxidant enzyme activities are important to tolerance to stress.

    Superoxide anion signalling at the beginning of stress is necessary for tolerance to stress.

    A slow Na + transport rate from roots to shoots can endow tolerance to stress.

  • Maize (Zea mays) is one of the world's three major food crops but sensitive to salinity at the seedling stage. Salinity/salt stress usually occurs due to gradually increased NaCl or under sudden exposure to NaCl, of which the latter is called salt shock (SS). However, little is known about physio-biochemical responses of maize to SS at the whole-plant level. The purpose of this study was to characterize the physio-biochemical response events of maize under SS. The experiments were conducted with four maize foundation parent inbred lines of Huangzao4, Chang7-2, Zheng58 and Ye478 under 150 mM NaCl for SS and after removal of SS in the nutrient solutions. The main findings were that the maize lines had no clear phase-order-response to SS, which suffered from the combined effects of osmotic stress, water deficiency, and Na+ accumulation-induced toxicity once SS occurred, and that SS-tolerant maize lines showed (1) timely increased activities of antioxidant enzymes (superoxide dismutase, peroxidase, catalase, and ascorbate peroxidase) and stronger superoxide anion radical-mediated signalling in roots at the beginning of SS, (2) a slow Na+ transport rate from roots to shoots especially in the early SS stage, and (3) opening of leaf stomata, and fine cell membrane integrity during SS. The related mechanisms of SS tolerance of maize were proposed and discussed.
    Graphical Abstract
  • Perennial grasses [e.g., switchgrass (Panicum virgatum), big bluestem (Andropogon gerardii), indiangrass (Sorghastrum nutans), little bluestem (Schizachyrium scoparium), Maasai love grass (Eragrostis superba), and bush ryegrass (Enteropogon macrostachyus)] are plant species that live for more than two years with deep root systems and the capacity to grow in a variety of climates[15]. Although often overlooked, perennial grasses serve an important role in ecosystems, particularly in maintaining soil health and biodiversity, climate change mitigation, and combating alien invasive plants (AIPs)[1,4]. Thus, they are simply natural allies for soil biodiversity conservation, invasive plant management, and climate change mitigation[6,7]. The deep root systems of perennial grasses help soil structure by improving aeration, increasing water infiltration, and lowering soil erosion[1,5]. Also, their extensive root network supports the stability of the soil, making it less susceptible to degradation and encouraging a healthier ecology overall[1,8]. Further, they play a key role in the nutrient cycle by maximizing nutrient utilization and minimizing leaching[9,10]. In addition, perennial grasses contribute organic matter to the soil through biomass, which decomposes over time and enriches the soil with critical nutrients[1113]. This process improves soil fertility, increasing productivity for other plant species, and agricultural activities[9,12].

    IAPs, also known as non–native or exotic species, are plants introduced to an ecosystem where they do not naturally occur[1416] and pose a severe ecological, economic, and social impacts[17,18]. Unlike native species, IAPs often lack natural enemies and diseases in their new environments, allowing them to proliferate unrestrictedly[19,20]. Their invasions lead to the displacement of native flora as they outcompete native species for resources i.e., light, water, and nutrients[21,22]. As a result, causing a reduction in biodiversity and the alteration of ecosystem functions, often forming dense monocultures that hinder the growth of other plants and disrupt habitats for native wildlife[23,24]. Moreover, IAPs can alter soil chemistry and hydrology thereby negatively impacting soil biodiversity[6,7,15,25]. IAPs can further impact human health by increasing allergens and providing a habitat for disease vectors[15]. Efforts to manage IAPs typically involve early detection, prevention, and rapid response, such as biological control, mechanical removal, and herbicide treatment[19,25,26]. Although the role of perennial grasses in combating IAPs has been seldom investigated, available studies show that effective management requires integrated eco-friendly management incorporating competitive native perennial grasses to suppress IAPs[6,8,15,27].

    Furthermore, perennial grasses are ecologically significant because they enhance species diversity and soil biodiversity i.e., living forms found in soil, which includes microorganisms (bacteria and fungi), mesofauna (nematodes and mites), and macrofauna, i.e., earthworms and insects[2832]. This diversity is critical to ecosystem function and plays an important role in nutrient cycling, soil structure maintenance, and plant growth promotion[29,30]. They contribute to nutrient-cycling activities by breaking down organic materials into simpler compounds that perennial grasses and other plants can consume, decomposing dead plants and animals, and releasing nutrients back into the soil, thus increasing soil fertility[3234]. Further, perennial grasses also promote plant-soil symbiotic relationships such as mycorrhizal associations and rhizobium symbioses, which improves soil health and plant growth[29]. These benefits are enhanced by perennial grasses' root exudates, which support both soil microbial diversity and activity, resulting in a more dynamic and resilient soil environment[1]. However, extreme weather events, such as floods and droughts, as well as IAPs can cause soil organism loss and structural damage, thereby impeding the roles of soil organisms[3537]. Further, increased temperatures can disrupt microbial activity and nitrogen cycling mechanisms, impacting soil health, and productivity[37,38]. Addressing these challenges needs long-term integrated management approaches that maintain natural ecosystems and increase soil biodiversity, as well as IAP control and climate change mitigation. For instance, promoting the use and maintaining the diversity of perennial grasses in rangelands and agricultural habitats[1,39,40].

    Climate change which is the average change in the earth's temperature and precipitation patterns can also disrupt the delicate balance of soil biodiversity[37,41]. It is driven primarily by human activities i.e., burning fossil fuels, deforestation, and industrial processes which lead to an unprecedented rise in greenhouse gases, such as carbon dioxide and methane in the atmosphere[37,42]. Often the earth's surface temperature increases concomitantly with these greenhouse gasses[41]. Increased temperatures contribute to sea-level rise, more frequent and intense heatwaves, wildfires, and droughts affecting biodiversity, water supply, and human health. Changes in precipitation patterns also lead to extreme weather events i.e., hurricanes, floods, and heavy rainfall, disrupting ecosystems and human societies[37]. It also negatively impacts biodiversity, as species must adapt, migrate, or face extinction due to altered habitats and shifting climate zones[36]. Addressing climate change requires global cooperation and robust policies aimed at reducing greenhouse gas emissions which include the use of eco-friendly approach, for instance, keeping the environment intact with native plants i.e., perennials grasses[43]. Perennial grasses (e.g., turfgrass) are considered potential for mitigating the effects of climate change because they have a high carbon sequestration capacity, storing carbon in both soil and aboveground biomass[4446]. They can contribute to reducing greenhouse gas levels by absorbing and storing carbon dioxide from the atmosphere in their roots and tissues, thus helping to mitigate climate change[44]. Furthermore, their capacity to minimize greenhouse gas emissions through reduced tillage and increased nitrogen use efficiency makes them an important component of habitat restoration to mitigate climate change impacts[43].

    Consequently, native perennial grasses have been recommended by various previous studies to be used for habitat restoration, including rangelands, because of their physiological and morphological traits, which have shown great potential to improve soil health and biodiversity, mitigate climate change, and combat IAPs[1,5,8,27,40,47]. By their competitive and morphological traits, several perennial native grass species found in African rangelands (e.g., African foxtail grass (Cenchrus ciliaris), horsetail grass (Chloris roxburghiana), rhodes grass (Chloris gayana), E. superba, and E. macrostachyus) and P. virgatum, S. nutans, S. scoparium, and A. gerardii in North America have been tested and recommended for ecological restoration[15].

    Preceding studies have demonstrated that perennial grasses have the potential to improve soil health and structure in rangelands and protected habitats[1,4850]. Unlike annual plants, which have shallow root systems, perennial grasses can penetrate deep into the soil, sometimes reaching depths of several meters as they have deep and extensive root systems[1,7,40]. These deep roots create channels that enhance soil aeration, allowing for better oxygen flow and water infiltration, thereby preventing soil compaction[49]. Perennial grasses contribute to soil stability by binding soil particles together, thereby preventing erosion (Fig. 1), which is important in ecosystems or habitats prone to heavy rainfall or wind[48,49]. This stabilization effect reduces the loss of topsoil, which contains the highest concentration of organic matter and nutrients essential for plant growth[44]. Moreover, perennial grasses have been reported to be efficient in nutrient cycling, a critical process for maintaining soil fertility[49]. For instance, their deep roots access nutrients in deeper soil layers, which might be unavailable to shallow-rooted plants[49,50]. These nutrients are then brought to the surface and incorporated into the plant biomass. When the grasses die back or shed leaves, these nutrients are returned to the soil surface as organic matter, making them accessible to other plants[32,49,51]

    Figure 1.  Diagram illustrating the multifaceted benefits of perennial grasses and their interconnected roles in promoting soil health, biodiversity, IAPs control, climate change mitigation, water retention, erosion control, and habitat provision. The arrows illustrate the complex interactions and synergies among these components, emphasizing the comprehensive ecological contributions of perennial grasses. The central position of perennial grasses highlights their pivotal role in these areas. This visual representation emphasizes how perennial grasses contribute to and enhance various aspects of ecosystem health and stability.

    Furthermore, perennial grasses enhance soil health and structure (Fig. 1), improving the soil's ability to retain water and withstand extreme weather events i.e., heavy rainfall and floods[44,49]. Their extensive root networks stabilize the soil, reducing erosion and runoff (Fig. 1), which are critical for maintaining soil fertility and agricultural productivity under variable climatic conditions[51]. The continuous growth and decay cycle of perennial grasses contributes to the slow but steady release of nutrients[52]. This slow release is beneficial for maintaining a stable nutrient supply, as opposed to the rapid nutrient depletion often seen in soils dominated by annual crops[50]. This process also helps in reducing nutrient leaching, where nutrients are washed away from the soil profile, particularly nitrogen, which is critical for plant growth[49]. Perennial grasses help to reduce N2O emissions; excess nutrients can lead to increased N2O emissions[10,11,53]. They also contribute significantly to the soil organic matter, which is a key component of soil health[52]. Organic matter consists of decomposed plant and animal residues, which improve soil structure, water retention, and nutrient availability[50,52]. The biomass produced by perennial grasses, both above and below ground, adds a substantial amount of organic material to the soil[52]. As the plant material decomposes, it forms humus, a stable form of organic matter that enhances soil structure by increasing its capacity to hold water and nutrients[52,54]. This is particularly important in dry regions e.g. in Africa, where water retention can be a limiting factor for crop growth[49]. The organic matter also provides a habitat and food source for a diverse array of soil organisms, including bacteria, fungi, and earthworms, which further contribute to soil fertility through their biological activities[43,52,54].

    Perennial grasses play a crucial role in enhancing soil biodiversity (abundance and diversity) and activities within the soil[31,32,51,54]. They provide critical habitats for soil fauna i.e., earthworms, nematodes, and arthropods (Fig. 1)[32,54]. Their complex root systems create a stable environment that supports a wide range of soil organisms[55]. Also, the root systems of perennial grasses exude a variety of organic compounds, including sugars, amino acids, and organic acids, which serve as food sources for soil biodiversity[54]. This continuous supply of root exudates and a stable environment fosters a diverse macro and microbial community, which is essential for maintaining soil health[31,43,54]. For instance, it was reported by Smith et al.[54] that in areas with abundant perennial grasses, a high soil macrofaunal biodiversity (i.e., Lumbricidae, Isopoda, and Staphylinidae) was observed. They further asserted that these grasses were beneficial to soil macrofauna as they increased the abundance and species diversity of staphylinid beetles, woodlice, and earthworms. In addition, Mathieu et al.[56] reported the influence of spatial patterns of perennial grasses on the abundance and diversity of soil macrofauna in Amazonian pastures. These findings suggest that well-managed perennial grasses are vital in enhancing soil macro and microbes in ecosystems[5456].

    These soil organisms perform various functions, including decomposing organic matter, fixing atmospheric nitrogen, and suppressing soil-borne diseases[29,30,32]. A diverse soil macro and microbial community can enhance nutrient cycling, making nutrients more available to plants[30,56]. Enhanced microbial diversity by perennial grasses contributes to the suppression of pathogens through competition and the production of antimicrobial compounds, thus promoting plant health[32]. They also help in maintaining soil structure, fertility, and overall ecosystem function[32]. For instance, earthworms, often referred to as 'ecosystem engineers', augment soil structure by creating burrows that improve aeration and water infiltration in perennial grass communities[31,51]. Their activity also helps mix organic matter into the soil, promoting nutrient cycling[31,32]. Nematodes and arthropods which feed on perennial grass species contribute to the decomposition process, breaking down organic matter and releasing nutrients that are vital for plant growth[31,54]. The presence of a diverse soil fauna community is indicative of a healthy soil ecosystem, which is more resilient to environmental stresses and disturbances[31].

    Furthermore, perennial grasses are considered as being instrumental in promoting plant-soil symbiotic relationships[43,54], which are crucial for plant health and soil fertility. One of the most well-known symbiotic relationships is between plants and mycorrhizal fungi[29,33]. These fungi colonize plant roots and extend their hyphae into the soil, increasing the root surface area and enhancing the plant's ability to absorb water and nutrients, particularly phosphorus. The relationship between perennial grasses and mycorrhizal fungi is mutually beneficial. The fungi receive carbohydrates produced by the plant through photosynthesis, while the plant gains improved access to soil nutrients and increased resistance to soil-borne pathogens[30]. This symbiotic relationship is particularly important in nutrient-poor soils, where mycorrhizal associations can significantly enhance plant growth and survival. Additionally, perennial grasses promote other beneficial plant-soil interactions, such as those involving nitrogen-fixing bacteria. These bacteria form nodules on the roots of certain perennial grasses, converting atmospheric nitrogen into a form that plants can use[29,30]. This process is essential for maintaining soil fertility, especially in ecosystems where nitrogen is a limiting nutrient.

    Perennial grasses are increasingly recognized for their role in climate change mitigation (Fig. 1)[43,44,57]. They can sequester carbon, reduce greenhouse gas emissions, and adaptation to climate variability[58,59]. Their deep root systems and grass-like characteristics make them highly effective in capturing and storing carbon[44]. These roots can penetrate deep into the soil and store carbon for extended periods[59]. Because of this, perennial grasses show potential to enhance the resilience of ecosystems to changing climatic conditions[44]. The roots of perennial grasses are more extensive and persistent compared to annual crops, allowing for greater carbon storage both in the root biomass and the soil[45,46,60]. This process of carbon sequestration involves capturing atmospheric carbon dioxide (CO2) through photosynthesis and storing it in perennial grass tissues (e.g., turfgrasses) and soil organic matter[4446]. Preceding studies have further shown that perennial grasses can sequester substantial amounts of carbon, contributing to the reduction of atmospheric CO2 levels[45,61]. In addition to carbon sequestration, perennial grasses can reduce greenhouse gas emissions through various mechanisms[43]. One of the primary ways is by reducing the need for frequent soil tillage, which is common in annual cropping systems. Tillage disrupts soil structure, releases stored carbon as CO2, and increases soil erosion[58,61]. Thus, with their long lifespan, perennial grasses can reduce the need for tillage, thereby minimizing CO2 emissions from soil disturbance[43,58].

    Moreover, perennial grasses can improve nitrogen use efficiency, reducing the need for synthetic fertilizers that are a major source of nitrous oxide (N2O) emissions—a potent greenhouse gas[53,62]. Their deep root systems enable them to access nutrients from deeper soil layers, reducing nutrient leaching and the subsequent emissions of N2O[53]. By optimizing nutrient use, perennial grasses contribute to lower greenhouse gas emissions associated with agricultural practices[63]. Also, perennial grasses are crucial for adapting to climate variability[44]. Their deep root systems allow them to access water from deeper soil layers, making them more resilient to drought conditions compared to annual crops[44]. This water use efficiency helps maintain plant growth and productivity even during periods of water scarcity, which are expected to become more frequent with climate change[49]. In general, perennial grasses support soil biodiversity conservation through habitat provision, climate change mitigation, and promoting ecosystem resilience[58]. Besides, these grasses are crucial for ecosystem stability and productivity, particularly in the face of climate change, and ensure the continued provision of ecosystem services (Fig. 1).

    Previous studies have shown that IAPs pose significant threats to ecosystems worldwide by displacing native species, altering habitats, and disrupting ecosystem functions and services[15,20,23,64]. Among the integrated management techniques to combat IAPs involves the use of competitive native plants (Fig. 1) such as perennial grasses[6,7,40]. These grasses, which live for more than two years with robust root systems, growth, and resilience to varying environmental conditions, offer several advantages in controlling IAPs[1,48]. Their competitive growth patterns and ability to restore and maintain native plant communities, and establish, and thrive in diverse habitats make them formidable competitors against invasive plants[1]. One of the primary ways perennial grasses combat IAPs is through competition for resources[48]. Their extensive root systems allow them to efficiently absorb water and nutrients, outcompeting IAPs that typically have shallower roots. This competitive edge limits the resources available to IAPs, inhibiting their growth and spread. For instance, species like P. virgatum and big A. gerardii are known for their deep roots, which can reach depths of up to 10 feet (3 m), providing them with a significant advantage over many IAPs[8,48]. They can also outcompete IAPs through their competitive growth patterns including quick establishment and forming dense canopies that shade out AIPs[1,8]. For example, native perennial grasses like S. nutans and S. scoparium have been shown to effectively compete with invasive species i.e., spotted knapweed (Centaurea stoebe) by limiting light availability and space for growth[8,48].

    Moreover, using their extensive root systems that stabilize the soil, perennial grasses can prevent erosion and invasions of IAPs[44]. Invasive plants i.e., carrot weed (Parthenium hysterophorus), cheatgrass (Bromus tectorum), and kudzu (Pueraria montana) can rapidly colonize disturbed soils, leading to severe erosion problems[20,65,66]. However, perennial grasses i.e., P. virgatum and big A. gerardii have been found to reduce erosion and creating an unfavorable environment for IAPs to establish owing to their deep fibrous root systems that hold the soil in place. Perennial grasses can also modify the microenvironment in ways that make it less conducive for IAPs[1,27,66]. They produce dense root mats that strengthen the organic matter content and soil structure, improving the fertility and health of the soil. The diversity and growth of native plant species is aided by improved soil conditions, which further promote biodiversity and inhibit IAPs by strengthening ecosystem resilience[48].

    Additionally, the use of perennial grasses in restoration has shown promising results in reclaiming areas overrun by IAPs and maintaining native plant communities that are disrupted by IAPs[8,66]. By planting a mix of native perennial grasses, land managers can restore ecological balance and prevent the re-establishment of IAPs[26]. These grasses provide long-term ground cover and habitat for wildlife, contributing to the overall health and stability of the ecosystem[1,8,54]. By reintroducing native perennial grasses into areas (e.g., rangelands and protected habitats) dominated by IAPs, ecosystems, and their biodiversity can be restored to their earlier conditions[27,39,67]. For instance, the use of native perennial grasses has been successful in restoring prairie ecosystems that were previously overrun by IAPs i.e., leafy spurge (Euphorbia esula) and purple loosestrife (Lythrum salicaria)[68]. Another important example of using perennial grasses to mitigate IAPs is the restoration of tallgrass prairies in the Midwest United States[8,66]. These prairies were historically dominated by native perennial grasses i.e., S. nutans and S. scoparium, however IAPs i.e., smooth brome (Bromus inermis) and reed canarygrass (Phalaris arundinacea) displaced them, leading to biodiversity loss and altered ecosystem functions[8,66,68]. Studies show that following the restoration of these invaded habitats using perennial grasses, native grasses successfully reestablished and reduced IAPs and promoting native biodiversity[66,67]. In addition, another notable example is the use of perennial grasses to restore riparian areas which were heavily invaded and impacted by IAPs i.e., giant reed (Arundo donax) and saltcedar (Tamarix spp.)[67,69]. Planting native perennial grasses like western wheatgrass (Pascopyrum smithii) and creeping wildrye (Elymus triticoides) in these areas helped to stabilize the soil, reduce erosion, and suppress IAPs, leading to improved riparian habitat quality and ecosystem resilience[18,66,67,69].

    Therefore, competitive suppressive perennial grasses are a crucial tool in the fight against IAPs and other weeds. Their competitive abilities, contributions to soil health, and role in ecosystem restoration makes them invaluable in managing and alleviating the impacts of IAPs. As research continues, the potential for perennial grasses to be integrated into broader IAP strategies remain significant, promising a more sustainable and ecologically sound approach to preserving native biodiversity.

    Perennial grasses are pivotal in enhancing soil biodiversity, mitigating climate change, and combating IAPs. Their deep root systems stabilize soils, support diverse soil faunal communities, and improve water retention. Besides, they are important grasses in sequestering carbon, reducing greenhouse gas emissions, suppressing IAPs, and supporting the reestablishment of native plant communities. Integrating perennial grasses into protected areas and rangelands management practices could offer a sustainable solution to pressing environmental challenges including invasions of IAPs. Stakeholders i.e., farmers, conservationists, ecologists, and land managers are advised to use perennial grass systems in their restoration practices, crop rotations, and pasturelands to enhance soil health and resilience. They are further commended to use perennial grasses for erosion control and to improve soil structure and fertility. Policymakers could develop and support policies that incentivize the use of perennial grasses in agricultural and restoration projects. Researchers, they are advised to conduct studies to quantify the long-term benefits of perennial grasses on soil biodiversity and climate change mitigation. Additionally, they can develop country or region-specific guidelines for the effective use of perennial grasses in different ecosystems. Hence, by integrating perennial grasses into our environmental stewardship strategies, we can ensure a thriving, balanced ecosystem capable of withstanding the impacts of climate change and IAPs.

    The author confirms sole responsibility for the following: review conception and design, and manuscript preparation.

    Data sharing not applicable to this article as no datasets were generated or analyzed during the current study.

    The author thanks all the colleagues who reviewed and proofread this article. This work was not supported by any funding agency.

  • The author declares that there is no conflict of interest.

  • [1]

    Roy SJ, Negrão S, Tester, M. 2014. Salt resistant crop plants. Current Opinion in Biotechnology 26:115−24

    doi: 10.1016/j.copbio.2013.12.004

    CrossRef   Google Scholar

    [2]

    Tuteja N. 2007. Mechanisms of high salinity tolerance in plants. Methods in Enzymology 428:419−38

    doi: 10.1016/S0076-6879(07)28024-3

    CrossRef   Google Scholar

    [3]

    Shavrukov Y. 2013. Salt stress or salt shock: which genes are we studying? Journal of Experimental Botany 64:119−27

    doi: 10.1093/jxb/ers316

    CrossRef   Google Scholar

    [4]

    Hasegawa PM, Bressan RA, Zhu JK, Bohnert HJ. 2000. Plant cellular and molecular responses to high salinity. Annual Review of Plant Physiology and Plant Molecular Biology 51:463−99

    doi: 10.1146/annurev.arplant.51.1.463

    CrossRef   Google Scholar

    [5]

    Ashraf M, Harris PJC. 2004. Potential biochemical indicators of salinity tolerance in plants. Plant Science 166:3−16

    doi: 10.1016/j.plantsci.2003.10.024

    CrossRef   Google Scholar

    [6]

    Parida AK, Das AB. 2005. Salt tolerance and salinity effects on plants: a review. Ecotoxicology and Environmental Safety 60:324−349

    doi: 10.1016/j.ecoenv.2004.06.010

    CrossRef   Google Scholar

    [7]

    Zhang H, Han B, Wang T, Chen S, Li H, et al. 2012. Mechanisms of plant salt response: insights from proteomics. Journal of Proteome Research 11:49−67

    doi: 10.1021/pr200861w

    CrossRef   Google Scholar

    [8]

    Deinlein U, Stephan AB, Horie T, Luo W, Xu G, et al. 2014. Plant salt-tolerance mechanisms. Trends in Plant Science 19:371−79

    doi: 10.1016/j.tplants.2014.02.001

    CrossRef   Google Scholar

    [9]

    Gupta B, Huang B. 2014. Mechanism of salinity tolerance in plants: physiological, biochemical, and molecular characterization. International Journal of Genomics 2014:701596

    doi: 10.1155/2014/701596

    CrossRef   Google Scholar

    [10]

    Athar HUR, Zulfiqar F, Moosa A, Ashraf M, Zafar ZU, et al. 2022. Siddique KHM. Salt stress proteins in plants: An overview. Frontiers in Plant Science 13:999058

    doi: 10.3389/fpls.2022.999058

    CrossRef   Google Scholar

    [11]

    Chauhan PK, Upadhyay SK, Tripathi M, Singh R, Krishna D, et al. 2022. Understanding the salinity stress on plant and developing sustainable management strategies mediated salt-tolerant plant growth-promoting rhizobacteria and CRISPR/Cas9. Biotechnology and Genetic Engineering Reviews 00:1−37

    doi: 10.1080/02648725.2022.2131958

    CrossRef   Google Scholar

    [12]

    Fu H, Yang Y. 2023. How plants tolerate salt stress. Current Issues in Molecular Biology 45:5914−34

    doi: 10.3390/cimb45070374

    CrossRef   Google Scholar

    [13]

    Ismail A, El-Sharkawy I, Sherif S. 2020. Salt stress signals on demand: Cellular events in the right context. International Journal of Molecular Sciences 21:3918

    doi: 10.3390/ijms21113918

    CrossRef   Google Scholar

    [14]

    Ma L, Liu X, Lv W, Yang Y. 2022. Molecular mechanisms of plant responses to salt stress. Frontiers in Plant Science 13:934877

    doi: 10.3389/fpls.2022.934877

    CrossRef   Google Scholar

    [15]

    Morton MJL, Awlia M, Al-Tamimi N, Saade S, Pailles Y, et al. 2019. Salt stress under the scalpel - dissecting the genetics of salt tolerance. The Plant Journal 97:148−63

    doi: 10.1111/tpj.14189

    CrossRef   Google Scholar

    [16]

    Zhou H, Shi H, Yang Y, Feng X, Chen X, et al. 2023. Insights into plant salt stress signaling and tolerance. Journal of Genetics and Genomics In press

    doi: 10.1016/j.jgg.2023.08.007

    CrossRef   Google Scholar

    [17]

    Abogadallah GM. 2010. Insights into the significance of antioxidative defense under salt stress. Plant Signaling & Behavior 5:369−74

    doi: 10.4161/psb.5.4.10873

    CrossRef   Google Scholar

    [18]

    Menezes-Benavente L, Kernodle SP, Margis-Pinheiro M, Scandalios JG. 2004. Salt induced antioxidant metabolism defenses in maize (Zea mays L.) seedlings. Redox Report 9:29−36

    doi: 10.1179/135100004225003888

    CrossRef   Google Scholar

    [19]

    Maas EV, Hoffman GJ, Chaba GD, Poss JA, Shannon MC. 1983. Salt sensitivity of corn at various growth stages. Irrigation Science 4:45−57

    doi: 10.1007/BF00285556

    CrossRef   Google Scholar

    [20]

    Alberico GJ, Cramer GR. 1993. Is the salt tolerance of maize related to sodium exclusion? I. Preliminary screening of seven cultivars Journal of Plant Nutrition 16:2289−303

    doi: 10.1080/01904169309364687

    CrossRef   Google Scholar

    [21]

    Fortmeier R, Schubert S. 1995. Salt tolerance of maize (Zea mays L.): the role of sodium exclusion. Plant, Cell & Environment 18:1041−47

    Google Scholar

    [22]

    Mansour MMF, Salama KHA, Ali FZM, Abou Hadid AF. 2005. Cell and plant responses to NaCl stress in Zea mays L. cultivars differing in salt tolerance. General and Applied Plant Physiology 31:29−41

    Google Scholar

    [23]

    de Azevedo Neto AD, Prisco JT, Enéas-Filho J, de Abreu CEB, Gomes-Filho E. 2006. Effect of salt stress on antioxidative enzymes and lipid peroxidation in leaves and roots of salt-tolerant and salt-sensitive maize genotypes. Environmental and Experimental Botany 56:87−94

    doi: 10.1016/j.envexpbot.2005.01.008

    CrossRef   Google Scholar

    [24]

    Azooz MM, Ismail AM, Abou Elhamd MF. 2009. Growth, lipid peroxidation and antioxidant enzyme activities as a selection criterion for the salt tolerance of three maize cultivars grown under salinity stress. International Journal of Agriculture and Biology 11:21−26

    Google Scholar

    [25]

    Schubert S, Neubert A, Schierholt A, Sümer A, Zörb C. 2009. Development of salt-resistant maize hybrids: The combination of physiological strategies using conventional breeding methods. Plant Science 177:196−202

    doi: 10.1016/j.plantsci.2009.05.011

    CrossRef   Google Scholar

    [26]

    Gondim FA, Gomes-Filho E, Costa JH, Mendes Alencar NL, Prisco JT. 2012. Catalase plays a key role in salt stress acclimation induced by hydrogen peroxide pretreatment in maize. Plant Physiology and Biochemistry 56:62−71

    doi: 10.1016/j.plaphy.2012.04.012

    CrossRef   Google Scholar

    [27]

    Parvaiz M. 2013. Response of maize to salt stress a critical review. International Journal of Healthcare Science 1:13−25

    Google Scholar

    [28]

    Pitann B, Mohamed AK, Neubert AB, Schubert S. 2013. Tonoplast Na+/H+ antiporters of newly developed maize (Zea mays) hybrids contribute to salt resistance during the second phase of salt stress. Journal of Plant Nutrition and Soil Science 176:148−56

    doi: 10.1002/jpln.201200597

    CrossRef   Google Scholar

    [29]

    Procházková D, Sairam RK, Lekshmy S, Wilhelmová N. 2013. Differential response of a maize hybrid and its parental lines to salinity stress. Czech Journal of Genetics and Plant Breeding 49:9−15

    doi: 10.17221/158/2011-CJGPB

    CrossRef   Google Scholar

    [30]

    Amdouni T, MrahS, Msilini N, Zaghdoud M, Ouerghiabidi Z, et al. 2014. Physiological and biochemical responses of two maize cultivars (Corralejo and Tlaltizapn) under salt stress. Journal of Stress Physiology & Biochemistry 10:246−58

    Google Scholar

    [31]

    Li Y, Wang TY. 2010. Germplasm base of maize breeding in China and formation of foundation parents. Journal of Maize Sciences 18:1−8

    doi: 10.13597/j.cnki.maize.science.2010.05.005

    CrossRef   Google Scholar

    [32]

    Ma D, Zhu J. 2012. Analysis on relationship between injuries and osmotic stress induced by salt stress in maize seedlings. Journal of Anhui Agricultural Sciences 40(34):16518−20

    doi: 10.13989/j.cnki.0517-6611.2012.34.128

    CrossRef   Google Scholar

    [33]

    Chen A. 1989. Observation of the states showing opening and closing stomata of the hybrid rice cultivar under the different temperatures with scanning electron microscope. Journal of Yuzhou University (Natural Sciences Edition) 6(3):45−49

    Google Scholar

    [34]

    Schützendübel A, Schwanz P, Teichmann T, Gross K, Langenfeld-Heyser R, et al. 2001. Cadmium-induced changes in antioxidative systems, hydrogen peroxide content, and differentiation in Scots pine roots. Plant Physiology 127:887−98

    doi: 10.1104/pp.010318

    CrossRef   Google Scholar

    [35]

    Giannopolitis CN, Ries SK. 1977. Superoxide dismutases: I. Occurrence in higher plants. Plant Physiology 59:309−14

    doi: 10.1104/pp.59.2.309

    CrossRef   Google Scholar

    [36]

    Beffa R, Martin HV, Pilet PE. 1990. In vitro oxidation of indoleacetic acid by soluble auxin-oxidases and peroxidases from maize roots. Plant Physiology 94:485−91

    doi: 10.1104/pp.94.2.485

    CrossRef   Google Scholar

    [37]

    Pine L, Hoffman PS, Malcolm GB, Benson RF, Keen MG. 1984. Determination of catalase, peroxidase, and superoxide dismutase within the genus Legionella. Journal of Clinical Microbiology 20:421−29

    doi: 10.1128/jcm.20.3.421-429.1984

    CrossRef   Google Scholar

    [38]

    Mishra NP, Mishra RK, Singhal GS. 1993. Changes in the activities of anti-oxidant enzymes during exposure of intact wheat leaves to strong visible light at different temperatures in the presence of protein synthesis inhibitors. Plant Physiology 102:903−10

    doi: 10.1104/pp.102.3.903

    CrossRef   Google Scholar

    [39]

    Verma S, Mishra SN. 2005. Putrescine alleviation of growth in salt stressed Brassica juncea by inducing antioxdative defense system. Journal of Plant Physiology 162:669−77

    doi: 10.1016/j.jplph.2004.08.008

    CrossRef   Google Scholar

    [40]

    Munns R, Schachtman DP, Condon AG. 1995. The significance of a two-phase growth response to salinity in wheat and barley. Australian Journal of Plant Physiology 22:561−69

    doi: 10.1071/pp9950561

    CrossRef   Google Scholar

    [41]

    García-Abellan JO, Egea I, Pineda B, Sanchez-Bel P, Belver A, et al. 2014. Heterologous expression of the yeast HAL5 gene in tomato enhances salt tolerance by reducing shoot Na+ accumulation in the long term. Physiologia Plantarum 152:700−13

    doi: 10.1111/ppl.12217

    CrossRef   Google Scholar

    [42]

    Jithesh MN, Prashanth SR, Sivaprakash KR, Parida AK. 2006. Antioxidative response mechanisms in halophytes: their role in stress defence. Journal of Genetics 85:237−54

    doi: 10.1007/BF02935340

    CrossRef   Google Scholar

    [43]

    Wang Y, Zhang W, Li K, Sun F, Han C, et al. 2008. Salt-induced plasticity of root hair development is caused by ion disequilibrium in Arabidopsis thaliana. Journal of Plant Research 121:87−96

    doi: 10.1007/s10265-007-0123-y

    CrossRef   Google Scholar

    [44]

    Sharma P, Jha AB, Dubey RS, Pessarakli M. 2012. Reactive oxygen species, oxidative damage, and antioxidative defense mechanism in plants under stressful conditions. Journal of Botany 2012:217037

    doi: 10.1155/2012/217037

    CrossRef   Google Scholar

    [45]

    You J, Chan Z. 2015. ROS regulation during abiotic stress responses in crop plants. Frontiers in Plant Science 6:1092

    doi: 10.3389/fpls.2015.01092

    CrossRef   Google Scholar

    [46]

    Shabala S, Cuin TA. 2008. Potassium transport and plant salt tolerance. Physiologia Plantarum 133:651−69

    doi: 10.1111/j.1399-3054.2007.01008.x

    CrossRef   Google Scholar

    [47]

    Cramer GR, Läuchli A, Polito VS. 1985. Displacement of Ca2+ by Na+ from the plasmalemma of root cells: A Primary response to salt stress? Plant Physiology 79:207−11

    doi: 10.1104/pp.79.1.207

    CrossRef   Google Scholar

    [48]

    Rawyler A, Arpagaus S, Braendle R. 2002. Impact of oxygen stress and energy availability on membrane stability of plant cells. Annals of Botany 90:499−507

    doi: 10.1093/aob/mcf126

    CrossRef   Google Scholar

    [49]

    Birben E, Sahiner UM, Sackesen C, Erzurum S, Kalayci O. 2012. Oxidative stress and antioxidant defense. World Allergy Organization Journal 5:9−19

    doi: 10.1097/WOX.0b013e3182439613

    CrossRef   Google Scholar

    [50]

    Cramer GR, Lynch J, Läuchli A, Epstein E. 1987. Influx of Na+, K+, and Ca2+ into roots of salt-stressed cotton seedlings: Effects of supplemental Ca2+. Plant Physiology 83:510−16

    doi: 10.1104/pp.83.3.510

    CrossRef   Google Scholar

    [51]

    Azaizeh H, Steudle E. 1991. Effects of salinity on water transport of excised maize (Zea mays L.) roots. Plant Physiology 97:1136−45

    doi: 10.1104/pp.97.3.1136

    CrossRef   Google Scholar

    [52]

    Lynch J, Läuchli A. 1988. Salinity affects intracellular calcium in corn root protoplasts. Plant Physiology 87:351−56

    doi: 10.1104/pp.87.2.351

    CrossRef   Google Scholar

    [53]

    Gilliham M, Dayod M, Hocking BJ, Xu B, Conn SJ, et al. 2011. Calcium delivery and storage in plant leaves: exploring the link with water flow. Journal of Experimental Botany 62:2233−50

    doi: 10.1093/jxb/err111

    CrossRef   Google Scholar

  • Cite this article

    Pan JL, Fan XW, Li YZ. 2023. Insights into physio-biochemical responses of maize to salt shock stress and removal of the stress at the whole-plant level. Tropical Plants 2:20 doi: 10.48130/TP-2023-0020
    Pan JL, Fan XW, Li YZ. 2023. Insights into physio-biochemical responses of maize to salt shock stress and removal of the stress at the whole-plant level. Tropical Plants 2:20 doi: 10.48130/TP-2023-0020

Figures(8)  /  Tables(1)

Article Metrics

Article views(2773) PDF downloads(384)

Other Articles By Authors

ARTICLE   Open Access    

Insights into physio-biochemical responses of maize to salt shock stress and removal of the stress at the whole-plant level

Tropical Plants  2 Article number: 20  (2023)  |  Cite this article

Abstract: Maize (Zea mays) is one of the world's three major food crops but sensitive to salinity at the seedling stage. Salinity/salt stress usually occurs due to gradually increased NaCl or under sudden exposure to NaCl, of which the latter is called salt shock (SS). However, little is known about physio-biochemical responses of maize to SS at the whole-plant level. The purpose of this study was to characterize the physio-biochemical response events of maize under SS. The experiments were conducted with four maize foundation parent inbred lines of Huangzao4, Chang7-2, Zheng58 and Ye478 under 150 mM NaCl for SS and after removal of SS in the nutrient solutions. The main findings were that the maize lines had no clear phase-order-response to SS, which suffered from the combined effects of osmotic stress, water deficiency, and Na+ accumulation-induced toxicity once SS occurred, and that SS-tolerant maize lines showed (1) timely increased activities of antioxidant enzymes (superoxide dismutase, peroxidase, catalase, and ascorbate peroxidase) and stronger superoxide anion radical-mediated signalling in roots at the beginning of SS, (2) a slow Na+ transport rate from roots to shoots especially in the early SS stage, and (3) opening of leaf stomata, and fine cell membrane integrity during SS. The related mechanisms of SS tolerance of maize were proposed and discussed.

    • Soil salinity is one of the major constraints to agricultural production[1] as many crop species are sensitive to salinity and generally cannot grow under NaCl at or over 100 mM[2], which is more likely to occur under an overlap of both drought and high temperature. The exposure to gradually increasing levels of NaCl is usually called salt stress, whereas sudden exposure is considered as salt shock (SS)[3]. The former often occurs in saline soils, and the latter rarely but occasionally takes place in cultivated lands that are vulnerable to flooding by large amounts of seawater[3].

      Studies on tolerance of plants to salt stress have been conducted across diverse plant species[1, 2, 49]. Even now, the mechanism of salt tolerance remains a hot topic in plant-related research fields, with numerous articles reviewing research progress[1016].

      It has been indicated that at the physiological level there are positive correlations between salt tolerance, activities of the antioxidant enzymes such as superoxide dismutase (SOD), peroxidase (POD), catalase (CAT) and ascorbate peroxidase (APX), and the synthesis of antioxidant compounds[8, 17].

      As one of the world's three major food crops, maize (Zea mays) is relatively sensitive to salt stress[18], more sensitive at emergence and seedling stages than at the flowering stage[19]. Maize planting has been expanded into salinity-affected lands because of the ever-growing demand for this crop, where this crop is bound to meet SS. Maize tolerance to salt stress is under intensive and extensive study[2030], but its response to SS, even for other plants, is still largely unclear.

      Huangzao4 (HZ4), Chang7-2 (C7-2), Ye478 (Y478) and Zheng58 (Z58) are important in-use foundation parent inbred lines for maize crossbreeding in China, of which C7-2 and Z58 are derivative inbred lines of HZ4 and Y478[31], respectively. Both HZ4 and C7-2 are of the Tangshan Sipingtou Chinese landrace germplasm, Y478 belongs to Reid's yellow dent germplasm introduced from modern American maize hybrids, and Z58 is from the Lvda red coda Chinese landrace germplasm[31]. According to our pre-experiments, HZ4, C7-2, Y478 and Z58 differed in SS tolerance. The hypothesis was that responses to SS would be different with maize lines differing in tolerance. In this study, we focused on how these maize inbred lines responded to SS in a 1× Hoagland nutrient solution supplemented with 150 NaCl.

    • Maize inbred lines of HZ4, C7-2, Y478 and Z58 were grown in a growth room that had a humidity of 60%−80%, temperatures of 28 °C (day) and 26 °C (night), and 12-h light of maximum light intensity of 13,000 lux provided by SYLVANIA Luxline Plus F58W/840 fluorescent light tubes (Germany).

      In brief, maize seeds were surface-sterilized with 75% ethanol and grown in sterile moist sand at 28 °C. At the two-leaf stage, the seedlings of health and uniform growth were treated by removal of the residual endosperm and then transplanted into holes of plastic foam boards, at least six holes and two seedlings per hole for each maize line under each treatment. The plastic foam boards were placed in square plastic pots containing 1× Hoagland nutrient solution at pH 6.0, where roots of the seedlings were completely immersed in the solution. During treatments, the nutrient solution was renewed once every 2 d and vigorously aerated for 15 min every 1 h. At the three-leaf stage, the nutrient solution in the pots was renewed with the nutrient solution supplemented with 150 mM NaCl for SS. The SS treatments were conducted for 5, 24, 48, and 72 h, respectively. The seedlings that were treated by SS for 72 h were transferred for removal of SS (RSS) into the new nutrient solution without the added NaCl and resumed growth for 48 h. Parallel control seedlings were those cultivated in the nutrient solution without the added NaCl.

    • Tissues were sampled from the fully expanded 2nd leaves and the roots at 10 a.m. The sampled tissues were immediately used, frozen in liquid nitrogen, or fixed for at least 24 h at 4 °C in the fixation solution containing 4% glutaraldehyde and 0.1 M of K2HPO4-KH2PO4 buffer (pH 7.2). The fixed tissues were used for the scanning electron microscopy (SEM) observation.

    • The RWC assay was performed as described in the literature[32] but with minor modifications. In brief, the sampled fresh leaves were immediately weighed (fresh weight, Wf), immersed in distilled water for 24 h, placed on dried filter paper to remove water of the leaf surface, and then weighed (saturated leaf fresh weight, Ws). The saturated leaves were further dried for 2 h at 105 °C and then for 7 h at 70 °C, and weighed (dry weight, Wd). The RWC was calculated as the following formula: RWC (%) = [(Wf-Wd)/(Ws-Wd)] × 100.

    • The leaves fixed in the fixation solution were washed three times with the KH2PO4-K2HPO4 buffer (pH 7.4) containing 4% (w/v) glutaraldehyde, and dehydrated for 30 min once sequentially in 30%, 50%, 70%, 80%, and 90% ethanol respectively, and then twice in 100% ethanol. The dehydrated leaves were then observed and imaged by using the Hitachi S-3400N SEM instrument following the procedures in the literature[33].

    • The root cell viability was evaluated following methods in the literature[34] but with minor modifications. Fragments (1 cm long) of fresh roots behind the root tips were stained for 30 min in 0.025% (w/v) Evans blue, rinsed for 15 min with deionized water, and then imaged.

      After imaging, roots were crushed with a glass rod, soaked for 30 min in 0.5 mL of a solution containing 50% (v/v) MeOH and 1% (w/v) SDS, heated for 15 min in water bath of 50 °C, and then centrifuged for 15 min at 14,000× g. The optical density value of the resulting supernatant at 600 nm was measured for estimation of Evans blue content by using the SHIMADZU UVmin-1240 spectrophotometer (Japan) and used to evaluate the cell viability.

    • The sample tissues were quickly rinsed with deionized water to remove the residues attached on the tissue surfaces, dried for 2 h at 105 °C, and then for 7 h at 70 °C until to a constant weight. The 0.1 g of the dried tissues was wet-ashed at 170 °C in 4 mL of concentrated sulfuric acid containing additional 5 drops of H2O2, and then analyzed by using the 6400 atomic absorption spectrophotometer (Shanghai Jinpeng Analytical Instruments Co., Ltd, China) following the manufacturer's instructions.

    • Two hundred mg of the frozen tissues were homogenized in 5 mL of a pre-chilled 50 mM NaH2PO4-Na2HPO4 buffer (pH 7.0) containing 1% polyvinylpyrrolidone (Guangdong Guanghua Chemical Factory Co. Ltd., Shantou, China), 1 mM ascorbic acid (Bio Basic Inc., Toronto, Canada) and 1 mM EDTA, and then centrifuged for 20 min at 15,000× g at 4 °C. The supernatant was used as the crude extract.

      SOD activity was assayed following the p-nitro blue tetrazolium chloride (NBT) method described in the literature[35]but with minor modifications. In brief, the photochemical reaction mixture was composed of 0.1 mL of the crude extract, 1.5 mL of 50 mM NaH2PO4-Na2HPO4 buffer (pH 7.0), 0.3 mL of 13 mM methionine, 750 µM NBT, 0.3 mL of 110 µM EDTA-Na2, 0.5 mL deionized water, and 10 µM riboflavin. The photochemical reaction was conducted for 20 min at 25 °C in a light incubator with 3,000 lux. The absorbance (A) value at 560 nm in the reaction was measured for estimation of SOD activity by using the spectrophotometer. SOD activity was estimated following the formula: = (ACK − AE) / (50% × ACK × Cpro × V), where ACK and AE were the A values of the control tubes and the reaction respectively, Cpro indicated the protein content in the crude extract (mg·L−1), and V represented the total crude extract used (mL).

      POD activity was assayed following the method in the literature[36]but with minor modifications. In brief, 0.02 mL of the crude extract reacted at 25 °C with 1 mL of 10 mM 3,3-dimethylglutaric acid-NaOH (pH 6.0) containing 5.5 mM guaiacol and 5.5 mM H2O2. The A470 nm value of the reaction was measured after reaction for 0, 30, 60, 90, 120 , and 180 s, by using the spectrophotometer. POD activity was estimated following the formula: = △A470 nm / (Cpro × V × t), where △A470 nm was the variation of the A value within the reaction time period, Cpro indicated the protein content in the crude extract (mg·L−1), V was the total crude extract used (mL), and t indicated the reaction time .

      CAT activity was assayed following the method in the literature[37] but with minor modifications. Briefly, 0.1 mL of the crude extract reacted at 25 °C with 1.4 mL of 0.05 M K2HPO4 (pH 7.0) containing 13.2 mM H2O2. The A240 nm value of the reaction was measured every 20 s for total 2 min by using the spectrophotometer. CAT activity was estimated following the formula: = △A 240 nm / (Cpro × V × t), where △A 240 nm was the variation of the A value within the reaction time period, Cpro indicated the protein content in the crude extract (mg·L−1), V was the total crude extract used (mL), and t indicated the reaction time.

      APX activity was measured following the ascorbate oxidation method[38]but with some modifications. The reaction was conducted at 25 °C in the 2-mL solution composed of 0.02 mL of the crude extract, 50 mM potassium phosphate (pH 7.0), 0.1 mM EDTA, 0.5 mM sodium ascorbate, and 0.1 mM H2O2. The A290 nm value of the reaction was measured after reaction for 0, 10, 20, 30, 40, 50, and 60 s by using the spectrophotometer. APX activity was estimated following the formula: = △A290 nm / (Cpro × V × t), where △A290 nm was the variation of the A value within the reaction time period, Cpro indicated the protein content in the crude extract (mg·L−1), V was the total crude extract (mL) used, and t indicated the reaction time.

    • The 0.1 mL of the crude extract and 0.1 mL of 0.6% thiobarbituric acid were mixed together, heated for 15 min in boiling water, immediately cooled on ice, and then centrifuged for 10 min at 1,698× g. The A value of the supernatant was measured at 532, 600, and 450 nm by using the spectrophotometer, respectively. The malondialdehyde content was calculated as a formula: = [6.45 × (A 532 nm – A 600nm) − 0.56 × A450 nm] / Cpro, where Cpro indicated the protein content in the crude extract (mg·L-1).

    • SAR content was estimated in accordance with the method in the literature[39]but with some modifications. First, 0.1 mL of the crude extract, 0.075 mL of 50 mM NaH2PO4-Na2HPO4 buffer (pH 7.8), and 0.025 mL of 10 mM hydroxylamine hydrochloride (Guangdong Guanghua Chemical Factory Co. Ltd., Shantou, China) were mixed and heated for 20 min at 25 °C. Then, 0.1 mL of 17 mM p-aminobenzene sulfonic acid (Bio Basic Inc., Toronto, Canada) and 0.1 mL of 7 mM α-naphthylamine (Shanghai Silian Chemical Co., Ltd., Shanghai, China) were added and allowed to further react for 30 min at 25 °C. The standard curve was plotted with the solution containing different concentrations of NaNO2, p-aminobenzene sulfonic acid, and α-naphthylamine. The A530 nm value of the reaction was measured by using the spectrophotometer and used to estimate SAR contents in the tissues as a formula: = (2 × X) / 20 × Vs × Cpro, where X was the standard curve reading, 2 was the dilution ratio of the used crude extract, 20 was the reaction time (min), Vs was solution sampled during the colour reaction (mL), and Cpro was the protein content of the crude extract (mg·L−1).

    • Statistical analyses of the data was conducted following the t test at a level of p < 0.05 using a programme in SPSS 13.0 software (www.spss.com).

    • Under control conditions, no significant differences were observed in phenotype among maize lines (Fig. 1a). The differences among maize lines in leaf phenotype occurred under SS and after RSS (Table 1; Fig. 1a). Under SS, Z58 showed no significant changes in leaf phenotype, and more than 95% of C7-2 leaves died after SS of 72 h (Table 1; Fig. 1a). After RSS, all seedlings of 72 h-SS-stressed Z58 survived, however, all seedlings of 72 h-SS-stressed C7-2 died (Table 1; Fig. 1a).

      Figure 1. 

      (a) Phenotype, (b) leaf RWC, and (c) root staining of maize inbred lines under SS and after RSS. The SS stress was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. In (b), the data were the means ± standard deviation (SD) of the fully expanded 2nd leaves of 5-leaf-old seedlings (n = 5−10) for each maize line under each treatment, and statistical analysis comparison was conducted between the same maize lines under control and the same SS stress, and between the same maize lines after SS of 72 h and after RSS. In (b), upper and lower cases of the same letter indicated a statistical significance at p < 0.05. In (c), fresh nodal roots (1 cm behind the root tip) from 5-leaf-old seedlings (n = 5−10) of each maize line were stained with Evans blue solution. The data of maize inbred line C7-2 after RSS were not available because of no surviving seedlings. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. RWC, Relative water content. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

      Table 1.  Phenotypes of seedlings of maize inbred lines under SS and after RSS.

      Maize lineYellowing of leavesDeath rate (%) of seedlings under SS forAfter RSS
      5 h24 h48 h72 hSurviving seedlings (%)
      Z58Slightly; The edge of about 10% of leaves after SS of 72 h0000100
      Y478Obviously; About 15% of leaves began turning yellow after SS of 48 h001.22.150.5
      C7-2Obviously; About 15% of the leaves began turning yellow after SS of 24 h0040.5950
      HZ4Somewhat like C7-20023.58048.65
      The SS stress was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. Leaves were observed and counted from leaves of 15−20 seedlings for each maize line. Survival rate after RSS referred to the percentag of surviving seedlings compared to seedlings stressed after SS of 72 h. C7-2, Maize inbred line Chang 7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

      Leaf RWC started to significantly decrease after SS of 5 h but had no significant difference among maize lines. The significant differences of maize lines in leaf RWC occurred after SS of 24, 48 and 72 h, respectively. Leaf RWC was highest in Z58 and lowest in C7-2 after SS of 72 h (Fig. 1b). Notably, leaf RWC of Z58 did not significantly fluctuate under SS (Fig. 1b).

      However, leaf RWC of Z58, Y478 and HZ4 significantly increased after RSS when compared to that in their respective maize lines SS-stressed for 72 h but it was still significantly lower than that of their respective control lines (Fig. 1b).

      The deeper the Evans blue staining indicated the less viability of cells. Consequently, the visible staining differences occurred among roots of maize lines after SS of 5 h and more significantly after SS of 24 h (Fig. 1c), where Evans blue-stained root zone was close to root tips of SS-stressed Z58 but relatively longer in SS-stressed lines of Y478, HZ4 and C7-2 (Fig. 1c).

      After RSS, Evans blue-staining was only at local root zone in SS-stressed Z58 but still in a longer root zone in SS-stressed lines of Y478 and HZ4. The staining depth of SS-stressed roots of maize lines followed Z58 < Y478 < HZ4 (Fig. 1c) but was shallower than that of their respective maize lines after SS of 72 h. Such staining differences were echoed partly by the quantitative assay of Evans blue (Fig. 2a).

      Figure 2. 

      Evans blue content in (a) fresh nodal roots, and Na+ content in (b) roots and (c) leaves of maize inbred lines under SS and after RSS. The SS was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. In (a), Evans blue content analysis was based on the Evans blue-stained fresh nodal roots (1 cm behind the root tip), where each datum was the mean ± SD from 5-leaf-old seedlings (n = 3−5) for each maize line under each treatment. In (b), each datum was the mean ± SD from a collection of roots of 5-leaf-old seedlings (n = 5−10). In (c), each datum was the mean ± SD from the fully expanded 2nd leaves of 5-leaf-old seedlings (n = 5−10). The statistical analysis comparison was conducted between the same maize line under control and the same SS stress, and between the same maize line after SS of 72 h and after RSS. The upper and lower cases of the same letter indicated a statistical significance at p < 0.05. The data of maize inbred line C7-2 after RSS were not available because of no surviving seedlings. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. SD, Standard deviation. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

      Both phenotype of shoots (Table 1; Fig. 1a) and root staining (Fig. 1c) under SS and after RSS indicated that SS tolerance degree of maize lines roughly followed Z58 > Y478 > HZ4 > C7-2.

    • Under control conditions, there were very slight but no significant differences in Na+ content of either roots (Fig. 2b) or leaves (Fig. 2c) among different maize lines.

      Na+ content in roots of SS-stressed maize lines significantly increased when compared to that in their respective control-treated lines, which roughly fluctuated as follows: highest in roots of Z58 after SS of 5 and 24 h, and no significant differences among maize lines after SS of 48 and 72 h (Fig. 2b). After RSS, Na+ content in roots of SS-stressed maize lines of Z58, Y478 and HZ4 significantly decreased when compared to that in their respective maize lines after SS of 72 h but it was still significantly higher than that in their respective control lines (Fig. 2a). It should be noted that after RSS Na+ content in roots of SS-stressed Z58 was still highest among SS-stressed maize lines, similar to the situation in its roots after SS of 5 and 24 h (Fig. 2b).

      Na+ content in leaves of SS-stressed C7-2 was highest among SS-stressed maize lines (Fig. 2c). After RSS (Fig. 2c), changes in Na+ content in leaves of SS-stressed maize lines were very similar to those in roots (Fig. 2b) of SS-stressed maize lines. Overall, the absolute Na+ content was much higher in leaves than in roots for each SS-stressed maize line.

    • Overall, K+ content in roots (Fig. 3a) and leaves (Fig. 3b) of Z58 was highest among maize lines under either control conditions or SS stress.

      Figure 3. 

      K+ content in (a) roots and (b) leaves, and Ca2+ content in (c) roots and (d) leaves of maize inbred lines under SS and after RSS. The SS was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. Each datum was the mean ± SD from the fully expanded 2nd leaves or a collection of roots of 5-leaf-old seedlings (n = 3−5) for each maize line under each treatment. The statistical analysis comparison was conducted between the same maize line under control and the same SS stress, and between the same maize line after SS of 72 h and after RSS. The upper and lower cases of the same letter indicated a statistical significance at p < 0.05. The data of maize inbred line C7-2 after RSS were not available because of no surviving seedlings. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. SD, Standard deviation. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

      As for K+ content in roots of SS-stressed maize lines, it showed no significant changes in Z58, significantly decreased in Y478 and HZ4, and significantly increased in C7-2 after SS of 5 h. K+ content tended to decrease although it fluctuated in some maize lines after SS of 24 h (Fig. 3a). After RSS, K+ content in SS-stressed maize lines of Z58 and Y478 was still lower than that in their respective control lines (Fig. 3a).

      Regarding K+ content in leaves of SS-stressed maize lines, it showed no change in Z58, significant decreases in Y478 and C7-2 and a significant increase in HZ4 after SS of 5 h when compared to that in their respective control lines. However, as the SS time prolonged, although K+ content was significantly lower than their respective control maize lines, it fluctuated obviously with maize lines. In general, K+ content in roots (Fig. 3a) and leaves (Fig. 3b) of C7-2 after SS of 48 and 72 h was lowest among SS-stressed maize lines. After RSS, K+ content in SS-stressed lines of Z58 and HZ4 was very close that in their respective control lines (Fig. 3b).

      With aspect to Ca2+ content in roots of SS-stressed maize lines, it significantly increased in Z58 and C7-2, and significantly decreased in Y478 and HZ4 after SS of 5 h (Fig. 3c). Ca2+ content in Z58 significantly decreased but remained relatively constant as SS exceeded 5 h. After RSS, Ca2+ content was still lower in SS-stressed Z58, sharply increased in SS-stressed Y478, and recovered to the control level in SS-stressed HZ4 when compared to that in their respective control maize lines (Fig. 3c).

      As regards Ca2+ content in leaves, it was highest in HZ4 and lowest in Z58 under control conditions. Interestingly, as for Ca2+ content under SS, it changed greatly with maize lines, either increased or decreased at some SS-time points. however, it was still lowest in SS-stressed Z58 when compred to that in teir respective control lines (Fig. 3d). After RSS, Ca2+ content significantly increased in SS-stressed lines of Z58 and Y478 (Fig. 3d).

    • In general, malondialdehyde content in roots (Fig. 4a) and leaves (Fig. 4b) of all SS-stressed maize lines tended to significantly increase as SS time prolonged when compared to that in their respective control maize lines, highest in roots (Fig. 4a) and higher in the most cases in leaves (Fig. 4b) of SS-stressed lines of HZ4 and C7-2. Notably, malondialdehyde content in roots (Fig. 4a) and (Fig. 4b) of SS-stressed Z58 and Y478 showed slight changes when SS time was over 24 h, not as dramatically increased as in other SS-stressed maize lines (Fig. 4a). After RSS, malondialdehyde content in roots (Fig. 4a) and leaves (Fig. 4b) of SS-stressed maize lines significantly decreased when compared to that in their respective lines that were SS-stressed for 72 h, but it was still higher than that in their respective control maize lines, highest in SS-stressed HZ4.

      Figure 4. 

      Malondialdehyde content in (a) roots and (b) leaves, and SAR content in (c) roots and (d) leaves of maize inbred lines under SS and after RSS. The SS was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. Each datum was the mean ± SD from the fully expanded 2nd leaves or a collection of the roots of 5-leaf-old seedlings (n = 3−5) for each maize line. The statistical analysis comparison was conducted between the same maize line under control and the same SS stress, and between the same maize line after SS of 72 h and after RSS. The upper and lower cases of the same letter indicated a statistical significance at p < 0.05. The data of maize inbred line C7-2 after RSS were not available because of no surviving seedlings. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. SAR, Superoxide anion radical. SD, Standard deviation. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

    • As for SAR production in roots, no significant differences were found among maize lines under control conditions (Fig. 4a). Under SS stress, SAR production significantly increased in maize lines but was highest in Z58 especially after SS of 5 h. However, SAR production tended to significantly decline in SS-stressed maize lines when SS time was at and over 24 h although it was higher than that in their respective control lines. After RSS, SAR production situation in SS-stressed lines of Z58, Y478 and HZ4 was almost the same as that in their respective maize lines that were SS-stressed for 72 h (Fig. 4c).

      Regarding SAR production in leaves, SAR production in Z58 and Y478 was much higher than that in HZ4 and C7-2 under control conditions (Fig. 4d). Under SS stress, SAR production was always much lower in most SS-stressed maize lines after SS of 5, 24, and 48 h, and significantly decreased in Y478 but increased in HZ4 and C7-2 after SS of 72 h (Fig. 4d) when compared to that in their respective control lines. After SS, SAR production was almost identical to that in their respective maize lines that were SS-stressed for 72 h (Fig. 4d).

    • As for SOD activity in roots, it showed differences among maize lines under control conditions, and significantly increased but was highest in Z58 under SS (Fig. 5a). For SOD activity in leaves, it was much higher in Z58 and Y478 under control conditions, and significantly decreased in Z58, Y478 and C7-2 but significantly increased in HZ4 after SS of 48 and 72 h (Fig. 5b) when compared to that in their respective control lines. After SS, SOD activity was higher in roots of SS-stressed maize lines (Fig. 5a) and only in leaves of SS-stressed HZ4 when compared to that in their respective control lines (Fig. 5b).

      Figure 5. 

      SOD activity in (a) roots and (b) leaves, and POD activity in (c) roots and (d) leaves of maize inbred lines under SS and after RSS. The SS was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. Each datum was the mean ± SD from the fully expanded 2nd leaves or a collection of the roots of 5-leaf-old seedlings (n = 3−5) for each maize line under each treatment. The statistical analysis comparison was conducted between the same maize line under control and the same SS stress, and between the same maize line after SS of 72 h and after RSS. The upper and lower cases of the same letter indicated a statistical significance at p < 0.05. The data of maize inbred line C7-2 after RSS were not available because of no surviving seedlings. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. POD, Peroxidase. RSS, Removal of SS. SD, Standard deviation. SOD, Superoxide dismutase. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

      Overall, the changing patterns of POD activity in either roots (Fig. 5c) or leaves (Fig. 5d) of maize lines either under control conditions and SS or after RSS were almost in line with those SOD activity in corresponding roots (Fig. 5a) or leaves (Fig. 5b) .

    • As for CAT activity in roots under SS, it significantly increased in all SS-stressed maize lines (Fig. 6a) but was much higher in SS-stressed Z58 than that in other SS-stressed maize lines especially after SS of 5 h. In leaves under SS, overall, CAT activity significantly increased after SS of 5, and 24 h for all SS-stressed maize lines and significantly decreased after SS of 48, and 72 h for SS-stressed maize lines of Z58 and Y478 (Fig. 6b). Notably, CAT activity was always higher in leaves of SS-stresed maize lines of HZ4 and C7-2 than that in their respective control lines (Fig. 6a). After SS, it was higher in roots (Fig. 6a) of all SS-stressed maize lines, and higher in leaves (Fig. 6b) of SS-stressed maize lines of Z58 and HZ4 than that in their respective control lines.

      Figure 6. 

      CAT activity in (a) roots and (b) leaves, and APX activity in (c) roots and (d) leaves of maize inbred lines under SS and after RSS. The SS was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. Each datum was the mean ± SD from the fully expanded 2nd leaves or a collection of the roots of 5-leaf-old seedlings (n = 3−5) for each maize line under each treatment. The statistical analysis comparison was conducted between the same maize line under control and the same SS stress, and between the same maize line after SS of 72 h and after RSS. The upper and lower cases of the same letter indicated a statistical significance at p < 0.05. The data were not available for maize inbred line C7-2 after RSS because of no surviving seedlings. APX, Ascorbate peroxidase; CAT, Catalase. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. SD, Standard deviation. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

      Overall, APX activity changes in roots (Fig. 6c) under control conditions and SS or after RSS were somewhat similar to CAT activity found in roots (Fig. 6a) although there were slight differences for some maize lines. As for APX activity in leaves, it was always higher in SS-stressed maize lines of HZ4 and C7-2, but significantly lower in SS-stressed maize lines of Z58 and Y478 after SS of 72 h when compared to that in their respective control lines (Fig. 6d). After RSS, APX activity was much higher in roots (Fig. 6c) of SS-stressed maize lines of Z58 and HZ4, and lower in leaves (Fig. 6d) of SS-stressed maize lines of Z58 and Y478 but much higher in leaves (Fig. 6d) of SS-stressed HZ4 when compared to that in their respective control lines.

    • The leaf stomata were always opened in Z58 under SS, began to close in HZ4 and C7-2 after SS of 24 h and in Y478 after SS of 48 h (Fig. 7a) when compared to those of their respective controls lines (Fig. 7b). After RSS, the leaf stomata were still opened in SS-stressed Z58, slightly opened in SS-stressed Y478, and still closed in SS-stressed HZ4 (Fig. 7a).

      Figure 7. 

      Stomatal behaviour in leaves of maize inbred lines under SS and after RSS. The SS was conducted with 150 mM NaCl. The RSS treatment was performed on maize plants stressed by SS for 72 h. The photos of the leaf stomata were taken by SEM from the central region of the front surface of the fully expanded 2nd leaves of 3-leaf-old seedlings (n = 5) for each maize line under each treatment. The data of maize inbred line C7-2 after RSS were not available because of no surviving seedlings. C7-2, Maize inbred line Chang7-2. HZ4, Maize inbred line Huangzao4. RSS, Removal of SS. SEM, Scanning electron microscopy. SS, Salt shock. Y478, Maize inbred line Ye478. Z58, Maize inbred line Zheng58.

    • In this study, responses of four maize inbred lines of Z58, Y478, HZ4 and C7-2 to SS and RSS were characterized. In terms of phenotype, SS tolerance was strongest for Z58 and weakest for C7-2 (Table 1; Fig. 1a).

      The decreased leaf RWC (Fig. 1b) and significantly increased Na+ content in roots (Fig. 2b) of SS-stressed maize lines after SS of 5 h suggest that maize suffers from the combined effects of water deficit, Na+ accumulation-induced osmotic stress at the whole-plant level once SS begins, somewhat differing from the two-phase ('osmotic' response/water deficit that dominates in Phase1 and the salt-specific response/salt toxicity in Phase 2) response model of plant growth under stepwise salt stress[40] .

      The salt tolerance mechanisms in plants partly depend on controlling Na+ uptake and transport from roots to shoots[6]. It was reported that maize cultivars of lower Na+ contents were more sensitive to salt than cultivars of higher Na+ contents[20, 27, 30]. The enhanced salt tolerance of tomato plants expressing yeast HAL5 gene was related to a lower Na+ transport rate from roots to shoots[41]. Interestingly, our results indicated that Na+ content showed massive differences in leaves (Fig. 2c) but not in roots (Fig. 2b) among SS-stressed maize lines, highest in C7-2 leaves (Fig. 2c). In addition, Na+ content increased highly in C7-2 leaves within a short SS time (viz. after SS of 5 h) but in leaves of other maize lines only after a longer SS time (viz. after SS of 72 h) (Fig. 2c). These results together with Evans blue staining of SS-stressed roots (Fig. 1c) suggest that Na+ transport rate from roots to shoots is maybe slower in SS-tolerant maize lines than in SS-sensitive maize lines, and further imply that maintaining Na+ homeostasis in cells of leaves is more important for SS tolerance of maize.

      A common oxidative stress on plants under salt stress results from over-production of reactive oxygen species (ROS) such as SARs[42]. High levels of ROS can damage cells[42]. However, an appropriate level of ROS is also important for plant growth and development because ROS plays a pivotal signalling role in stress-triggered tolerance mechanisms[4345]. Therefore, a balance between production and removal of ROS must be tightly regulated to tolerate stress[17, 44]. In this study, SAR levels were overall much higher in roots than in leaves of SS-stressed maize especially after SS of 5 h (Fig. 4c). Z58, the most tolerant maize line, had the highest SAR level in roots (Fig. 4c) and the lowest Na+ content in leaves after SS of 5 h (Fig. 2c). However, C7-2, the most sensitive maize lines, had a lower SAR level in roots after SS of 5 h (Fig. 4c) and the highest Na+ level in leaves after SS of 72 h (Fig. 2c). These results strongly indicate that SS-tolerant maize lines can easily generate SAR signalling in roots than SS-sensitive maize lines at the onset of SS, and further implicate that the Na+-induced SAR signalling is probably involved in mediating the Na+ transport from roots to shoots and/or in balancing intracellular Na+.

      The intracellular K+/Na+ ratio is a key determining trait of salt tolerance[46]. Lower K+ levels can further increase Na+ toxicity under salt stress because Na+ can compete with K+ for enzyme activation and protein biosynthesis[46]. Coupling of the lowest K+ contents (Fig. 3b) with the highest Na+ contents (Fig. 2c) in C7-2 leaves after SS of 5, 48, and 72 h suggest that Na+ accumulation under SS likely leads to leakiness of more cytosolic K+ in SS-sensitive maize than in SS-tolerant maize, in agreement with the prior viewpoints[47].

      The peroxidation of the cell membrane by ROS is one of the main causes of membrane damage, resulting in production of malondialdehyde[48, 49]. The high malondialdehyde contents in both roots (Fig. 4a) and leaves (Fig. 4b) in SS-stressed maize lines of HZ4 and C7-2 suggest that keeping the cell membrane stable is of great importance in maize tolerance to SS.

      High Ca2+ levels benefit plants under salt stress by compensating/minimizing the Na+-induced leakiness of cytosolic K+[50], increasing the relative availability of water for maize growth[51], and maintaining K+/Na+ selectivity[46]. Salt stress can cause a decrease in Ca2+ influx and an increase in Ca2+ efflux from the maize root cells[52]. During the first phase after approximately 2-3 weeks of salt stress applied in a hydroponic nutrient solution via daily NaCl increases, salt-sensitive maize cultivar 8023 had higher concentrations of Ca2+ than did salt-tolerant maize cultivar Pioneer 3906, although Ca2+ concentrations in shoots decreased in both two cultivars[21]. In cotton treated by SS in a 0.1× modified Hoagland solution supplemented with NaCl and CaCl2, Ca2+ influx increased in proportion to salt concentration (ranging from 150 to 250 mM NaCl)[50]. In this study, although Ca2+ content fluctuated greatly among roots of maize lines during SS (Fig. 3c), it gradually increased in leaves of Z58 and Y478 as SS time prolonged, and significantly decreased in leaves of HZ4 and C7-2 after SS of more than 24 h (Fig. 3d). Such discrepancies among different studies may be due to differences in treatment conditions/processes and materials. Anyway, our results suggest that maintaining high levels of Ca2+ in leaves is important to enhance maize tolerance to SS. These results together strongly indicate that Ca+/K+/Na+ balance is most important for palnt tolerance to salt stress but differ with plant species.

      SOD, POD, CAT and APX are major antioxidant enzymes for plants to cope with oxidative damage under abiotic stresses[9, 17, 42, 43]. In this study, that the activities of the enzymes in SS-stressed maize increased but were highest activities in roots of Z58 in a short SS time (viz. after SS of 5 h) (Figs 5 & 6) suggest that increasing the activities of the antioxidant enzymes in roots is more significant at the initial SS phase than the late SS phase for maize to tolerate SS. This is likely because roots are only one tissue that is directly exposed to SS environments, on the other hand, the early and timely increase in the enzyme activity is conducive to the reconstruction of the antioxidant systems for maize to adapt to the ensuing SS. In addition, the differences in the enzyme activities between roots and leaves and among different maize lines under SS (Figs 5 & 6) implies that the utilization of antioxidant systems under SS varies with tissues and maize lines, with APX activity pattern as an example which significantly decreased in leaves of Z58 but significantly increased in leaves of HZ4 and C7-2 under SS (Fig. 6d).

      The opening and closing of leaf stomata affect the entrance of CO2 into leaves, of which the stomatal closure not only causes the accumulation of ROS[42, 53] but also inhibits the production of osmoprotectants and radical scavengers[6]. The leaf stomata were always opened in Z58 and closed earlier in other maize lines under SS (Fig. 7a). The more significantly increased SAR levels in leaves of Y478, HZ4 and C7-2 as SS time prolonged (Fig. 4d). These results suggest that opening of the leaf stomata is particularly crucial to enhance maize tolerance to SS.

      Taken all results together, the related mechanisms of SS tolerance of maize as well as a possible way to improve maize SS tolerance by spraying Ca and K fertilizer were proposed (Fig. 8).

      Figure 8. 

      Schematic mechanisms of maize tolerance to SS, and a possible measure to improve tolerance to SS. A possible measure to improve maize tolerance to SS by spraying Ca and K fertilizer onto leaf surfaces was suggested and shown. APX, Ascorbate peroxidase; CAT, Catalase; POD, Peroxidase; ROS, Reactive oxygen species; SAR, Superoxide anion radical; SOD, Superoxide dismutase; SS, Salt shock.

    • Maize has no clear processes of phase-order-response to SS, which suffers from the combined effects of osmotic stress, water deficiency, and Na+ accumulation-induced toxicity once SS occurs. Stronger tolerance of maize to SS is characterized by (1) timely increases in activities of antioxidant enzymes (SOD, POD, CAT and APX) and a stronger SAR-mediated signalling necessary to trigger the relevant tolerance mechanisms in roots once SS occurs; (2) a slow Na+ transport rate from roots to shoots especially in the early SS stage; and (3) opening of leaf stomata, and fine cell membrane integrity to prevent leakage of Ca2+ and K+ under SS. However, these mechanisms should be verified with more maize lines in future.

    • The authors confirm contribution to the paper as follows: study conception and design: Li YZ, Fan XW; data collection: Pan JL; analysis and interpretation of results: Li YZ, Pan JL; draft manuscript preparation: Li YZ. All authors reviewed the results and approved the final version of the manuscript.

    • All data generated or analyzed during this study are included in this published article.

    • We are grateful to Professors Yu Li and Yun-Su Shi, the Institute of Crop Sciences, CAAS, who kindly supplied the maize seeds.

      • The authors declare that they have no conflict of interest.

      • Received 28 April 2023; Accepted 17 October 2023; Published online 7 November 2023

      • Timely increased antioxidant enzyme activities are important to tolerance to stress.

        Superoxide anion signalling at the beginning of stress is necessary for tolerance to stress.

        A slow Na + transport rate from roots to shoots can endow tolerance to stress.

      • Copyright: © 2023 by the author(s). Published by Maximum Academic Press on behalf of Hainan University. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (8)  Table (1) References (53)
  • About this article
    Cite this article
    Pan JL, Fan XW, Li YZ. 2023. Insights into physio-biochemical responses of maize to salt shock stress and removal of the stress at the whole-plant level. Tropical Plants 2:20 doi: 10.48130/TP-2023-0020
    Pan JL, Fan XW, Li YZ. 2023. Insights into physio-biochemical responses of maize to salt shock stress and removal of the stress at the whole-plant level. Tropical Plants 2:20 doi: 10.48130/TP-2023-0020

Catalog

  • About this article

/

DownLoad:  Full-Size Img  PowerPoint
Return
Return