Plant Physiology Preview. Published on February 16, 2016, as DOI:10.1104/pp.15.02026
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Short title: Singlet oxygen and vacuolar cell death
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Corresponding Author:
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Robert Fluhr
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Address:
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Department of Plant and Environmental Sciences
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Weizmann institute of science
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Rehovot, 76100
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Israel
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Telephone:
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+972-8-9342175
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Email:
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[email protected]
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Research Area:
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Signaling and Response
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Title: Singlet oxygen induced membrane disruption and serpin-protease balance in
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vacuolar driven cell death in Arabidopsis thaliana
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Authors:
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Eugene Koh1, Raanan Carmieli2, Avishai Mor1 and Robert Fluhr1
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Department of Plant and Environmental Sciences, Weizmann Institute, Rehovot, Israel
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Department of Chemical Research Support, Weizmann Institute, Rehovot, Israel
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One sentence summary:
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ROS such as singlet oxygen can affect vacuolar membrane integrity bringing about cell
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death that is further modulated by the balance of lytic proteases and their cognate
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inhibitors.
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Footnotes
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Funding information:
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We wish to acknowledge the support of the I-CORE Program of the Planning and
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Budgeting Committee and The Israel Science Foundation (757/12) and the Lerner Family
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Plant Science Research Fund and the Israel Science Foundation (grant No. 1596/15).
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Corresponding author:
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[email protected]
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Abstract
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Singlet oxygen plays a role in cellular stress either by providing direct toxicity or through
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signaling to initiate death programs. It was therefore of interest to examine cell death, as
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occurs in Arabidopsis thaliana, due to differentially localized singlet oxygen
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photosensitizers. The photosensitizers rose bengal and acridine orange were localized to
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the plasmalemma and vacuole, respectively. Their photoactivation led to cell death as
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measured by ion leakage. Cell death could be inhibited by the singlet oxygen scavenger
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histidine in treatments with acridine orange but not with rose bengal. In the case of
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acridine orange treatment, the vacuolar membrane was observed to disintegrate.
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Concomitantly, a complex was formed between a vacuolar cell-death protease,
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RESPONSIVE TO DESSICATION 21 (RD21) and its cognate cytoplasmic protease
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inhibitor ATSERPIN1. In the case of rose bengal treatment, the tonoplast remained intact
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and no complex was formed. Overexpression of AtSerpin1 repressed cell death, only
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under acridine orange photodynamic treatment. Interestingly, acute water stress showed
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accumulation of singlet oxygen as determined by fluorescence of Singlet Oxygen Sensor
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Green, by electron paramagnetic resonance (EPR) spectroscopy and the induction of
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singlet oxygen marker genes. Cell death by acute water stress was inhibited by the singlet
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oxygen scavenger histidine and was accompanied by vacuolar collapse and the
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appearance of serpin-protease complex. Overexpression of AtSerpin1 also attenuated cell
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death under this mode of cell stress. Thus, acute water stress damage shows parallels to
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vacuole-mediated cell death where the generation of singlet oxygen may play a role.
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Introduction:
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Singlet oxygen is a reactive oxygen species (ROS) that can be produced in a light
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dependent manner in the photosynthetic apparatus of plants or accumulate during certain
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biochemical reactions. It is a strong electrophile, and can undergo several types of
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reactions with biological molecules. Of note are cycloaddition reactions on double bonds,
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forming hydroperoxides via the ene reaction; and endoperoxides via the Diels-Alder
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reaction. It is also able to react with thiols, leading to the production of sulfoxides and
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sulfinic and sulfonic acids (Triantaphylides and Havaux, 2009).
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Singlet oxygen is a strong oxidant of polyunsaturated fatty acids (PUFAs) and is
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the main source of photooxidative damage in chlorophyll containing cells in Arabidopsis
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(Triantaphyllides et al., 2008). The resultant peroxidation reaction initiated by singlet
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oxygen can create a kink in the hydrophobic fatty acid tail causing irregularities in the
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membrane ultrastructure and vesicular budding (Riske et al., 2009); or cross linking and
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aggregation of membrane proteins (Kukreja and Hess, 1992; Beaton et al., 1995). Hence,
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damage to membrane function by singlet oxygen might arise from two distinct
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mechanisms; one through a purely physical mechanism of membrane disruption, and the
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other through failure of membrane pumps which are critical components in the
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maintenance of homeostasis.
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Cellular detoxification of singlet oxygen can be via physical energy transfer or
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through chemical reactions. The former type involves energy exchange between singlet
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oxygen and molecules like carotenoids. The latter type of chemical quenching can occur
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between singlet oxygen and ascorbate, where the quencher is oxidized and consumed.
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Recycling of the quencher can occur through enzymatic means, such as the ascorbate-
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glutathione pathway, or via de novo synthesis (Triantaphylides and Havaux, 2009). Other
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examples of dissipation occur through specific interaction with amino acids e.g. histidine
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to form endoperoxides (Nilsson et al., 1972; Matheson et al., 1975; Ishibashi et al, 1996).
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In addition to direct cytotoxicity, singlet oxygen possesses a signaling function
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(Kim et al., 2008). For example, in the dark, FLUORESCENT (flu) mutants accumulate
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excess protochlorophyllide (Pchlide) in the chloroplasts. Pchlide, a precursor of
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chlorophyll (Meskauskiene et al., 2001) acts as a photosensitizer and generates singlet
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oxygen, which leads to cell death in Arabidopsis plants when shifted from dark to light
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conditions (op den Camp et al., 2003). The light-generated cell death and gene induction
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in flu plants depend on EXECUTER1 and 2 implying that singlet oxygen plays a
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signaling role rather than direct toxicity (Wagner et al., 2004; Lee et al., 2007). However,
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other stresses that stimulate the generation of singlet oxygen in the chloroplast, e.g. high-
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light in the chlorina 1 mutant, were found to be independent of EXECUTER genes and
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may indicate direct toxicity (Ramel et al., 2013).
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Recent work has emphasized light-independent singlet oxygen production in
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response to wounding or the addition of elicitor in the dark (Mor et al., 2014).
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Additionally, transcriptome signatures characteristic of singlet oxygen were found to be
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pervasive in many biotic and abiotic stress conditions. Organelles such as the nucleus,
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mitochondria and peroxisome have been reported to be potential sources of singlet
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oxygen in root tips (Mor et al., 2014). In animal systems, singlet oxygen was shown to be
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involved in both signaling and membrane photo-oxidation in the cell (Klotz et al., 2003)
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and in phagocytic immune response (Steinbeck et al., 1992). Hence, the role of singlet
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oxygen in signaling and cell death is ubiquitous and not restricted to either light
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dependence or chlorophyll containing cells.
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Plant cell vacuoles are regulators of intracellular and organismal homeostasis and
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serve as cellular waste recovery stations, where misfolded proteins and damaged
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organelles are delivered for degradation and disposal. Vacuolar proteases that are
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important in protein and degradome processing include VACUOLAR PROCESSING
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ENZYME (VPE) and RESPONSIVE TO DESSICATION-21 (RD21) (Hara-Nishimura
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et al., 2005; Gu et al., 2012). These proteases can also carry out death-promoting
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functions in plant programmed cell death (PCD) (Hatsugai et al., 2004; Kuriyama and
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Fukuda, 2002; Hara-Nishimura et al., 2005; Lampl et al., 2013). It has been proposed that
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vacuolar mediated cell death could proceed by two distinct mechanisms (Hara-Nishimura
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and Hatsugai, 2011); the destructive pathway, involving disruption of the tonoplast, and a
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non-destructive pathway, in which the tonoplast membrane remains intact. The non-
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destructive way is proteasome dependent, and facilitates the secretion of anti-pathogenic
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substances from the vacuole into the apoplast through specific pores. The destructive
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pathway shares many similar features with necrotic type PCD pathways in plants such as
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the hypersensitive response (HR) (Hatsugai et al., 2004), and the formation of tracheary
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elements (TE) in xylem development (Kuriyama, 1999).
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When the vacuolar membrane is disrupted, eg. during elicitor induced vacuolar
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cell death, proteases like RD21 escape into the cytoplasm, thus exacerbating cell death
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(Lampl et al., 2013). Its cognate protease inhibitor, AtSerpin1, serves as a potential
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protective cell death check-point by acting as a suicide substrate, thus inactivating the
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protease. In one model, when such inhibitors are in excess, cells protect themselves from
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intracellular damage and survive, but when proteases exceed their inhibitors, the surplus
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proteases are able to degrade cellular proteins, leading to cell death. Thus, it has been
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suggested that serpins and proteases exist in a balance governed by their stoichiometric
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amount and the integrity of compartment membranes (Lampl et al., 2013). Singlet
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oxygen, if targeted towards membranes of hydrolytic organelles such as the vacuole,
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could result in the direct disruption of membranes, unleashing their stored proteolytic
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activity. Here, we demonstrate that singlet oxygen produced in the vacuole, as well as the
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balance of proteases and their cognate inhibitors, can be components of vacuole-mediated
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cell death. We further demonstrate that acute water stress in the dark can lead to cell
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death in a manner that partially mimics singlet oxygen-driven vacuolar type cell death.
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Results
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Localization of photodynamic singlet oxygen generators and cellular damage
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With the aim of better understanding singlet oxygen toxicity and its possible
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relationship to vacuole-mediated cell death we chose to investigate 2 different
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photosensitizers, rose bengal (RB) and acridine orange (AO). RB and AO were applied to
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5 day old Arabidopsis seedlings in the dark (Figure 1A). The plant tonoplast marker γ-
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TONOPLAST INTRINSIC PROTEIN (TIP) aquaporin fused to the cyan fluorescence
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protein (CFP) marker was used to assist in determining their localization (Nelson et al.,
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2007). In the case of RB, staining was observed in the cell periphery, whereas AO was
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localized to the vacuole. When a hypertonic solution of 1 M mannitol was applied, the
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vacuole shrunk revealing RB stain mainly in the plasmalemma (Figure 1B, top and
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Supplementary Figure S1). When seedlings treated with AO were subjected to the same
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treatment, the vast majority of AO could be detected within the confines of the vacuole
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while a residual amount was observed in the cell wall (green color; Figure 1A and B;
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bottom). RB has been shown previously to localize to the cell periphery in Arabidopsis
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cell cultures and to the chloroplast (Gutierrez et al., 2014). AO is sequestered in acidic
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compartments like the vacuole where it becomes protonated and trapped in the cation
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form to which the tonoplast membrane is impermeable (Oparka et al., 1991; Boller et al.,
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1986).
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Singlet oxygen is among the major reactive oxygen species shown to be involved
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in chloroplast-based lipid peroxidation in oxidative-type cell death e.g. in high light stress
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(Triantaphylides et al., 2008). It is therefore of interest to examine the ramifications of its
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accumulation in other cellular compartments. Both AO and RB treated plants showed
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light-dependent cell death as determined by ion leakage (Figure 2A). The kinetics of the
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cell death profiles of the 2 photosensitizers appear to be different. RB treatment induced
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rapid light-dependent ion leakage as early as 2 h but decelerated cell death towards the
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later time points (Figure 2B). In contrast, AO treated samples showed a delayed cell
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death profile, which started to accelerate after 24 h (Figure 2C).
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Histidine scavenges singlet oxygen and hydroxyl radicals with a high degree of
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specificity by both physical and chemical mechanisms (Nilsson et al., 1972; Matheson et
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al., 1975; Ishibashi et al, 1996). Histidine effectively scavenged RB induced singlet
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oxygen production in vitro as shown by measuring the changes in fluorescence of the
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singlet oxygen probe, SOSG, after light treatment (Supplementary Figure S2). We tested
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the scavenging effect of histidine against both RB and AO generated phototoxicity in
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vivo. AO photosensitized cell death was reduced by histidine (Figure 2E). However, RB
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treated samples showed an unexpected increase in apparent cell death in the presence of
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histidine
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intermediates (Cadet and Di Mascio, 2006) that can be toxic. Apparently, the local
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cellular milieu can impact on the efficacy of histidine to act as a protective scavenger.
(Figure
2D).
Photo-oxidized
histidine
forms
reactive
endoperoxide
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AO but not RB induces vacuolar destruction
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Cell death rates approaching 40% were observed after 72 h light treatment in both
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RB and AO applications (Figure 2B & C). A residual fluorescent signal from AO was
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evident after 72 h in the light and dark that differed from the initial distribution (compare
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Figures 1A and 3A). In addition, in the light the vacuolar membrane marker was
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disrupted (Figure 3A, compare light and dark). Additional CFP fluorescent material of
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unknown origin also appears to accumulate in the vacuole after extended 72 h of AO
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treatment in the dark (Figure 3A). In contrast, in RB-treated tissue after 72 h, the residual
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fluorescent signal was mostly bleached in the light but the vacuolar membrane appeared
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intact (Figure 3B, compare light and dark). Sytox Green staining was used to examine the
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integrity of membranes in RB treatments. Sytox Green enters cells with compromised
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plasmalemma and binds to nucleic acids. Sytox Green staining of RB treated tissue
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showed that the cell outer membrane was rendered permeable by RB phototoxicity in the
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light but not in the dark, with little effect on the tonoplast structure (Supplementary
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Figure S3, compare light and dark). Thus, photodynamic cell death with RB and AO
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showed distinctive kinetics of ion leakage and differential effect of the photodynamic
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process on the tonoplast membrane, suggesting distinct mechanisms of cell death.
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AO but not RB induces mixing of cytoplasm and vacuolar compartments
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The ablation of the tonoplast membrane by AO treatment in the light suggested
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the possibility that application of AO could mimic vacuolar-mediated cell death that is
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characterized by the mixing of cytoplasmic and vacuolar contents. To visualize this at the
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molecular level we took advantage of the fact that the AtSerpin1-HA protease inhibitor is
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expressed in the cytoplasm and can form a covalent complex with the vacuolar RD21
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protease upon mixing of cellular compartments (Lampl et al, 2010). Photodynamic cell
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death was induced in an AtSerpin1-HA overexpression line by treating plants with or
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without 100 μM RB or AO under continuous light conditions. Extracts were treated with
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cysteine protease inhibitor E-64 to prevent further formation of complex (Roberts et al.,
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2011) and fractionated on non-reducing SDS-PAGE gel (Figure 4). Inspection of the
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Coomassie stained gel shows a gradual decrease in the levels of the large subunit of
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RIBULOSE-1,5-BISPHOSPHATE CARBOXYLASE/OXYGENASE (RUBISCO) that
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is accelerated in AO and RB. The decrease in RUBISCO may indicate a more rapid
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turnover of cellular proteins due to cell death over 72 h. Immunoblots were developed
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with AtSerpin1 antibody to detect serpin-protease complex. Significantly more complex
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accumulated in the samples treated by AO compared to the control with concomitant
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disappearance of the free serpin at 45 kD. The increased appearance of cleaved 'spent
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serpin' likely represents turnover of complex that frees the residual N-terminal (37 kD)
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fragment. An additional higher molecular weight immune-reactive polypeptide was
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present in AO-treated samples in the light and the dark (Figure 4; Supplementary Figure
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S4). This may be due to the presence of an additional AO-induced serpin-protease
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complex. The presence of additional serpin-protease complexes has been noted before
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(Lampl et al., 2010). The presence of this putative higher molecular weight complex in
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the dark and the light indicates that it is likely not due to the production of singlet
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oxygen. In the RB treated sample no complex or free serpin were detected although spent
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serpin was observed. Spent serpin can result from disassociation of complex or activation
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of non-cognate proteases that cleave the serpin (Lampl et al., 2010). In the dark, control,
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AO and RB treatments are similar except for the presence of the higher molecular weight
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immune-reactive polypeptide in the AO treated sample mentioned above (Supplementary
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Figure S4). All samples showed a gradual accumulation of complex but to a lesser extent
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than observed in the light with AO and may be a result of autophagic cellular processes
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that utilize RD21 protease and are stimulated under dark starvation conditions. Taken
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together, the enhanced formation of complex implies that AO photodynamic damage led
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to mixing of cytoplasmic and vacuolar contents. In contrast, the photodynamic activity of
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RB led to cell death without apparent vacuolar collapse or mixing of compartments.
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AtSerpin1 in the cytoplasm can partially protect cells from vacuolar-mediated cell death
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Serpins are localized to the cytoplasm and are thought to serve a cytoprotective
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function from vacuolar cell death proteases (Lampl et al., 2013). We asked if
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photodynamic cell death could bring about the release of vacuolar proteases that would
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be impacted by the presence of serpins in the cytoplasm. To this end, the response of wild
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type (WT), overexpression AtSerpin1-HA (HA), AtSerpin1 knockout (AtSerpin1-ko) and
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RD21 knockout (RD21-ko) plant lines were analyzed (Figure 5). AtSerpin1-HA lines
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showed a protective effect to photodynamic mediated damage following AO treatment
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(Figure 5B, p-value < 0.001). The RD21-ko lines were less effective in the reduction of
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cell death (p-value < 0.057), whilst the AtSerpin-ko lines were not statistically different
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from the wild type lines. This may be due to gene redundancy. Under dark conditions no
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statistical difference can be measured between any of the lines (Supplementary Figure
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S5). Significantly, none of the RB treated plant lines showed differences from each other
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in the light (Figure 5A). This indicates that the serpin-protease system does not play a
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role in RB-motivated cell death and is consistent with the lack of formation of serpin-
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protease complex (Figure 4). Taken together, the results suggest that serpin
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overexpression or the absence of RD21 protease can mitigate vacuolar-type cell death. It
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is possible that the presence of multiple proteases in the vacuole explain why only partial
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protection was afforded by RD21-ko and AtSerpin1-HA lines.
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Dehydration treatment in the dark produces singlet oxygen and vacuolar disruption
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Transcriptomic signatures for singlet oxygen appear in many diverse stresses
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including water stress (Mor et al., 2014). It is therefore of interest to compare tissue death
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induced by this stress with photodynamic-induced cell death. Leaf tissue was dehydrated
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for various times (1-3 h) and then analyzed over 72 h. Dehydration treatments of 2 and 3
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h led to 20% and 60% cell death respectively after 72 h. Cell death was rapid and reached
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a plateau 2 h after treatment (Figure 6A). The application of histidine decreased damage
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by dehydration as measured by ion leakage by more than 30% (Figure 6C, p < 0.01). In
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order to further substantiate a possible role for singlet oxygen in this type of tissue death,
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Arabidopsis seedlings were dehydrated in the dark and treated with the highly specific
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Singlet Oxygen Sensor Green (SOSG) probe that forms diagnostic endoperoxides in the
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presence of singlet oxygen (Flors et al., 2006). A rise of fluorescence, indicating singlet
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oxygen-dependent formation of endoperoxide, was detected rapidly in the cytoplasm
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after 15 min of dehydration (Figure 6B). After 30 min of drying treatment in the dark, the
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SOSG reporter was observed to permeate the cell. In contrast, in wet control samples
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placed in the dark for similar times, no SOSG fluorescence was detected and the
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tonoplast remained intact (Supplementary Figure S6).
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To
quantitate
singlet
oxygen
production,
the
spin
trap,
4-hydroxy-
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tetramethylpiperidine (4-hydroxy-TEMP) was employed. It shows enhanced specificity
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for singlet oxygen that can be detected by electron paramagnetic resonance spectroscopy
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(EPR) (Nakamura et al., 2011). EPR spectra obtained from RB and dehydration treated
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plants indicate a comparable production of singlet oxygen. However, in AO treated plants
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a reliable spectrum was not observed (Figure 6D, Supplementary Figure S7). AO
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treatment may generate a signal that is too low for detection or the singlet oxygen is
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sequestered from the spin trap probe. As singlet oxygen will react with membrane lipids
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to form hydroperoxyl lipid radicals and lipid peroxidation, these products can be detected
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by the Thiobarbituric Acid Reactive Substances (TBARS) assay (Hodges et al., 1999). As
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shown in Figure 7, RB, AO and dehydration treatments led to the detection of statistically
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significant increases in TBARS equivalents. Taken together the evidence from SOSG
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fluorescence, 4-hydroxy-TEMP oxidation and the detection of lipid peroxidation products
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by TBARS are consistent with singlet oxygen production leading to a degree of lipid
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peroxidation.
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The rapid permeation of the vacuole by SOSG may indicate a contribution of
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vacuolar collapse to dehydration-induced cell death. To examine this, tonoplast integrity
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was followed by serpin-protease complex formation. The amount of AtSerpin1-protease
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complex and spent cleaved serpin increased as a result of dehydration stress in mature
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plants (Figure 8A) or in seedlings (Supplementary Figures S8 and S9). This observation
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indicates a degree of mixing of cellular compartments due to dehydration stress but to a
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lower degree than observed for AO treatment. To test this observation further, wild type,
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overexpression AtSerpin-HA, AtSerpin1-ko and RD21-ko were examined for their
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sensitivity to drought simulation. In this case, only a small but statistically significant
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reduction (Figure 8B; 18%; p < 0.05) in ion leakage was noted in the overexpression
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AtSerpin1-HA line. Taken together, the results indicate that acute water stress induced a
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degree of permeabilization of the vacuolar membrane as indicated by the distribution of
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SOSG fluorescence, disruption of the CFP tonoplast marker and by the formation of
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serpin-protease complex.
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Gene expression markers for singlet oxygen
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To further characterize the types of ROS produced under photodynamic and
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dehydration stress, qRT-PCR analysis was carried out with gene sets specific for types of
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ROS. Singlet oxygen was shown to induce gene sets At3g61190, At5g64870 and
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At3g01830 whereas superoxide/hydrogen peroxide induced At5g01600, At4g21870 and
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At1g71030 (op den Camp et al., 2003). As expected RB and AO showed specific
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induction of the singlet oxygen probes, whilst methyl viologen (MV) preferentially
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induced the expression of the superoxide/hydrogen peroxide probes (Figure 9A). The
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strength of gene induction by AO appears to be less then RB; a result that is consistent
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with its reduced quantitative EPR and TBARS measurements. Strikingly, the gene
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induction pattern of dehydration treatment is robust and closely follows that of singlet
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oxygen (Figure 9B).
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Discussion
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Singlet oxygen generators that localized to the plasma membrane and vacuole
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respectively, generate very different patterns of cell death. RB accumulated in the cell
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periphery and stimulated rapid ion leakage. The permeabilization of the plasmalemma by
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RB treatment left the tonoplast intact, as evidenced by the entry of Sytox Green, but the
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integrity of the tonoplast marker was maintained. RB also generated singlet oxygen that
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was measurable by singlet oxygen spin traps as well as by lipid peroxidation, established
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by TBARS assay. In contrast, AO was sequestered in the vacuole and stimulated slower
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ion leakage without detectable singlet oxygen production by EPR. Trivially one may
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suspect that no singlet oxygen was made, however this is inconsistent with the detection
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of lipid peroxidation by the TBARS assay and the finding that both treatments resulted in
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the induction of singlet oxygen specific genes. The discrepancy can be reconciled by
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noting that AO-generated singlet oxygen is less amenable to detection by spin-trap probes
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(Ito, 1978; Decuyper et al., 1984). In addition, the vacuole may be less accessible to the
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spin trap probe.
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AO treatment was characterized by extensive tonoplast collapse and the release of
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vacuolar cell death proteases that could be followed by formation of serpin-protease
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complexes. This occurred even though the amount of singlet oxygen produced by AO is
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low indicating that the vacuolar membrane is sensitive to this level of singlet oxygen.
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When the vacuole membrane is ruptured and the amount of released protease exceeds the
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capacity of the serpin, the subsequent widespread proteolysis of intracellular proteins
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disrupts cellular homeostasis (Yamashita, 2004). This stimulates a positive feedback loop
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that spirals to cell death that can be partially reversed in AtSerpin1 overexpression lines
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due to formation of inactive serpin-protease complexes. Significantly, photodynamic cell
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death as rendered by AO shows a degree of commonality with vacuolar mediated
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programmed cell death as described for certain plant-pathogen interactions (Hara-
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Nishimura and Hatsugai, 2011; Dickman and Fluhr 2013).
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In an analogous manner, AO applied to malignant melanoma cells was shown to
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accumulate in cellular acidic compartments that were disrupted in a photodynamic
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process that led to cell death (Hiruma et al., 2007). Death by RB is different and is likely
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through plasmalemma membrane permeabilization that initiates rapid ion leakage.
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Indeed, cercosporin, a lipophilic singlet oxygen generator secreted by necrotrophic
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phytopathogens of the Cercospora species is thought to accumulate in the membranes of
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host cells (Daub and Briggs, 1983; Daub and Ehrenshaft, 2000). Rapid electrolytic
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leakage was detected from a variety of plant leaf discs treated with cercosporin within
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minutes of exposure to light (Daub and Briggs, 1983; Macri and Vianello, 1979). Thus,
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RB and AO initiate qualitatively different pathways of cell death and each partially
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mimics distinct physiological situations.
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The role of singlet oxygen in promoting cell death is an open question. The direct
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oxidative action of singlet oxygen, especially on lipids and membrane bound proteins
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suggests that singlet oxygen can exert its toxicity through modulating membrane integrity
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(Riske et al., 2009; Kukreja and Hess; 1992; Beaton et al., 1995). Alternatively, singlet
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oxygen produced by the flu mutant in the chloroplast has been shown to induce cell death
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through signaling components rather than by direct toxicity. The expression of flu-
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induced stress genes requires the genes EXECUTER1&2 (Wagner et al, 2004; Lee et al.,
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2007; Kim et al., 2012; Kim and Apel, 2013). Additionally, under high light stress the
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photodynamic overflow caused by singlet oxygen circumvents the EXECUTER1&2
369
signaling pathway to directly induce cell death (Kim et al., 2012).
370
Dehydration stress was shown here to directly involve singlet oxygen
371
accumulation based on SOSG fluorescence and EPR measurements. These observations
372
are consistent with attenuation of cell death by the scavenger histidine and by induction
373
of singlet oxygen-specific genes. While tonoplast disruption was prominent during acute
374
water stress, the formation of serpin-protease complex and protection by AtSerpin1
375
overexpression was less robust compared to photodynamic vacuolar destruction using
376
AO. This may be due to the rapidity of death or alternatively, acute water stress cell death
377
may induce a conglomerate of destructive forces of which vacuolar integrity is but one
378
layer.
379
It is thought that drought accelerates cell death through photooxidative stress. In
380
one mechanism, stomatal closure that limits water loss also restricts CO2 intake, leading
381
to a reduction of the net photosynthetic rate that causes the overall generation of ROS
382
(Mittler, 2002; de Carvalho, 2008). The main ROS that are produced in this fashion are
383
O2-, H2O2 and singlet oxygen through the over reduction of the light harvesting
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15
384
photosystems and photorespiration (de Carvalho, 2008; Krieger et al., 2008). However as
385
shown here, and previously for root tissue (Mor et al., 2014), drying of plant tissue in the
386
dark also leads to ROS production in the form of singlet oxygen.
387
The source of singlet oxygen in driving chloroplast-induced cell death in the light
388
is chlorophyll. The source of singlet oxygen in the dark is unknown. Singlet oxygen has a
389
short half-life implying that its production is cell autonomous. Also, the presence of
390
singlet oxygen in the dark in the cytoplasm of leaf epithelial cells that are largely devoid
391
of chloroplasts indicate that chlorophyll is not a source. Evidence was recently presented
392
for singlet oxygen production in other organelles such as the mitochondria, peroxisomes
393
and in general in membrane rich regions (Mor et al., 2014). Singlet oxygen could arise in
394
the dark by reaction of O2- and H2O2 via the Haber-Weiss reaction (Khan and Kasha,
395
1994). For example, singlet oxygen lipid peroxidation signatures were present when
396
methyl viologen (MV), which is an inducer of O2- and H2O2, was applied to Arabidopsis
397
leaf discs (Triantaphyllides et al., 2008). However, the results here show that acute water
398
stress induces marker genes related to singlet oxygen rather than O2- or H2O2. Singlet
399
oxygen could also arise through the Russell mechanism in the chemical reactions
400
produced by peroxylipid radicals that are initiated by lipoxygenase activity (Kanofsky
401
and Axelrod, 1986; Miyamoto et al., 2007). Indeed, drought activated increases in
402
lipoxygenases were detected in Arabidopsis leaves (Gigon et al., 2004), and roots
403
(Grebner et al., 2013). A ubiquitous stress signaling mechanism for activation of
404
lipoxygenases may involve cellular calcium. Calcium is rapidly elevated during drought
405
stress (Knight et al., 1997) and was also shown to activate both lipoxygenase activity
406
(Nelson et al., 1977) and the general quasi-lipoxygenase activity of hemoproteins (Iwase
407
et al., 1998). It will be of interest to see how singlet oxygen integrates into the temporal
408
sequence of events from stress to ROS generation to changes in gene expression (Laloi
409
and Havaux, 2015; Dietz, 2014).
410
Production of singlet oxygen that is independent of light and chloroplasts appears
411
to be a common component in vacuolar cell death. Whether singlet oxygen directly
412
causes membrane permeabilization, or by working through some other means, remains to
413
be elucidated. Singlet oxygen transcriptome signatures are also manifest in other abiotic
414
and biotic stresses (Mor et al., 2014). Hence, it would be important to characterize the
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16
415
role of singlet oxygen in these stresses, and examine if there are similarities between the
416
resulting types of cell death.
417
418
419
Materials and Methods
420
421
Plant growth conditions and treatments
422
Arabidopsis thaliana (ecotype Columbia-0) seedlings (5 days to 3 weeks) were
423
grown under white light in a 16 h light / 8 h dark cycle at 21°C on Murashige and Skoog
424
medium, supplemented with 1% sucrose and 0.8% (w/v) phytoagar (Invitrogen).
425
Arabidopsis plants (8-12 weeks) were grown in soil under a 12 h light/dark cycle at 21°C.
426
All the in vivo experiments treated with RB and AO were supplemented with 1% sucrose,
427
whilst all the dehydration experiments were performed in ddH2O. For the in vitro singlet
428
oxygen assay 10 μM of RB (Sigma) and 1 μM of SOSG (Sigma) were prepared in ddH2O
429
and fluorescence measured at 485 nm excitation and 525 nm emission after exposing to
430
white light (30 μE) for 5 min intervals over 45 min.
431
432
Cell death assays with ion leakage
433
Ion leakage was performed on ten 5 day old seedlings or 5 leaf discs (6 mm; 8-12
434
weeks) per well in a 12 well plate (Corning) in 5 ml of ddH2O or 1% sucrose as stated,
435
with 3 biological replicates for each treatment. The samples were equilibrated in the light
436
for 1 h, and solutions replaced. Following this step, if photosensitizers (RB and AO) were
437
used, incubation was performed in the dark for 1 h, followed by 3 washes. For drought
438
treatment, leaf discs or seedlings were placed on dry Whatman paper in the dark for time
439
lengths as stated. Ion conductance was measured by sampling 100 μl using a Horiba B-
440
173 compact conductivity meter. If histidine was used, solutions were made in ddH2O or
441
1% sucrose as stated, adjusted to pH 7.2, and added after the wash step but before
442
baseline measurement. Samples were incubated under white light (30 μE) or in the dark
443
(wrapped in aluminum foil) as indicated. Maximum ion leakage was obtained after
444
boiling the samples in the microwave for 30 s, followed by another 15 s of boiling, and
445
then cooling at room temperature for 2 h. Relative cell death was calculated by
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17
446
subtracting the baseline and obtaining the ratio over the maximum ion leakage. Means
447
and standard errors were obtained thereafter.
448
449
Immunoblots
450
Extraction of proteins and their fractionation on reducing and non-reducing
451
denaturing gels has been described in Roberts et al., (2011). Membranes were developed
452
with anti-AtSerpin1 antibodies (1:1000) and secondary anti-guinea pig horseradish
453
peroxidase (1:3000) or with anti-RD21 antibodies (1:1000), and secondary anti-mouse
454
horseradish peroxidase (1:3000).
455
456
Confocal microscopy
457
Confocal microscopy analysis was carried out on 5-8 day old tonoplast-labeled
458
CFP marker line (vac-ck CS16256; Nelson, et al 2007). All images were taken with either
459
an Olympus IX81 FV1000D Spectral Type, or a Nikon A1 confocal microscope. For
460
CFP, excitation was at 405 nm using an argon laser and emission was at 440 nm. SOSG
461
staining was performed by incubation with 100 μM SOSG diluted in ddH2O in the dark
462
for 20 mins. Sytox Green staining was performed by incubation with 250 nM Sytox
463
Green diluted in ddH2O in the dark for 20 mins. For staining by Sytox Green, Singlet
464
Oxygen Sensor Green (SOSG, Invitrogen) and AO, excitation was at 488 nm and
465
emission was at 525 nm. For RB staining, excitation was at 561 nm and emission was at
466
580 nm. The various photosensitizer and drought treatments were performed on 5-8 day
467
old transgenic stable marker lines. Seedlings were equilibrated in ddH2O or 1% sucrose
468
as stated for 1 h in the light, and solutions replaced. Following this step, if
469
photosensitizers (RB and AO) were used, incubation was performed in the dark for 1h,
470
followed by 3 wash steps, and exposed to light for the time lengths stated. For drought
471
treatment, seedlings were placed on dry Whatman paper in the dark for time lengths as
472
stated, and stained with SOSG as indicated.
473
474
RNA Extraction and qRT-PCR Analysis
475
Arabidopsis seedlings (2-week-old, 7 seedlings per biological replicate, 3
476
replicates) were used for each treatment. RNA was extracted from frozen tissues using a
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18
477
standard Trizol extraction method (Sigma-Aldrich). DNase I (Sigma-Aldrich)-treated
478
RNA was reverse transcribed using a high-capacity complementary DNA reverse
479
transcription kit according to the manufacturer’s instructions (Quanta Biosciences). For
480
qRT-PCR analysis, the SYBR Green method (KAPA Biosystems) was used on a Step
481
One Plus platform (Applied Biosystems) with a standard fast program. qRT-PCR primers
482
were designed in Snapgene software. All qRT-PCR primer sequences are listed in
483
Supplemental Table S1.
484
485
EPR Spectroscopy and Sample Preparation
486
Arabidopsis seedlings (30 seedlings, 2-week-old) were used for each treatment.
487
The seedlings were equilibrated in ddH2O for dehydration or 1% sucrose for AO and RB
488
in ambient light for 2 h, replacing the medium after 1 h. All further incubation and
489
extraction procedures were carried out in the dark. For dehydration treatment, seedlings
490
were incubated with 0.4 M of 2,2,6,6-Tetramethyl-4-piperidinol (4-hydroxy-TEMP,
491
Sigma-Aldrich) for 30 min, and washed six times with ddH2O. Samples were then placed
492
on dry Whatman paper for 0, 30, 60, 90 min. Wet samples were floated in a petri dish
493
with ddH2O for similar times as a control. For RB and AO treatments, 100 μM of RB or
494
AO was co-incubated with 0.4 M 4-hydroxy-TEMP prepared in 1% sucrose for 30 min,
495
washed and placed in the light (30 μE) or dark for 2 h. Samples were harvested by flash
496
freezing in liquid nitrogen and were homogenized in a shaker using glass beads. The
497
samples were then dissolved in 1 ml of 0.2 M potassium phosphate buffer, pH 7.4,
498
containing 10 mM EDTA and 20 mM N-ethyl-maleimide, and then incubated in a rotator
499
for 15 min at 4°C. The extracts were then centrifuged at 14,000 rpm at 4 °C for 20 min,
500
and the supernatant was used for the EPR measurements. EPR measurements were
501
acquired at room temperature with a Bruker ELEXYS E500 spectrometer operating at X-
502
band frequencies (9.5 GHz) and a 100 kHz modulation frequency. The spectra exhibit a
503
typical TEMPO triplet EPR spectra (aN = 15.8 G) and signal intensity values of the third
504
peak of the nitroxide spectrum were obtained by calculating the difference between the
505
maximum and minimum points, and normalized to the wet weight of the sample and the
506
volume of extraction buffer used. Measurements of 4-hydroxy-TEMPO production
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19
507
induced by singlet oxygen were calculated by plotting against a standard curve of 4-
508
hydroxy-TEMPO.
509
510
TBARS assay
511
Arabidopsis seedlings (2-week-old, 5 seedlings per biological replicate, with 3 replicates)
512
were used for each treatment. The seedlings were equilibrated in distilled, deionized
513
water (ddH2O) or 1% sucrose in ambient light for 2 h, replacing the medium with fresh
514
ddH2O or 1% sucrose after 1 h. For RB and AO treatments, samples were incubated in
515
the dark with 100 μM RB or AO in 1% sucrose for 1 h, washed and then incubated in the
516
light (30 μE) or dark for various time points as stated. For dehydration treatment, samples
517
were incubated on dry Whatman paper or ddH2O in the dark for the various time points
518
stated. Samples were harvested by flash freezing in liquid nitrogen and were
519
homogenized in a shaker using glass beads. TBARS assay was performed according to
520
the method of Hodges et al., 1999. The samples were pre-weighed before treatment, and
521
TBARS values were normalized according to the wet weights.
522
523
Acknowledgments
524
We wish to acknowledge the support of the Lerner Family Plant Science Research Fund
525
and the I-CORE Program of the Planning and Budgeting Committee and The Israel
526
Science Foundation (757/12) and the Israel Science Foundation (grant No. 1596/15).
527
528
List of author contributions
529
EK formulated the hypotheses and carried out the experimental work and wrote the
530
manuscript; RC, AM and RF formulated the hypotheses and wrote the manuscript.
531
532
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20
533
Figure legends
534
Figure 1. Confocal microscopy of Arabidopsis seedlings showing localization of the
535
photosensitizers RB and AO. A, Recombinant tonoplast marker line (5 day old, CFP)
536
treated with 100 μM RB (Top) and AO (Bottom) respectively. B, Seedlings (5 day old,
537
CFP) treated with 100 μM RB (Top) and AO (Bottom) respectively, and mounted on a
538
microscope slide with 1M mannitol to allow for cell plasmolysis. Seedlings were
539
observed by confocal microscopy as described in the Material and Methods. Scale bar 20
540
μm.
541
542
Figure 2. Ion leakage in Arabidopsis plants showing effects of photosensitizers AO and
543
RB in the light and dark. A, Ion leakage assay. Leaf discs were treated with 300 μM AO
544
and RB respectively and incubated for 0, 24, 48, 72 h in the light (L) or dark (D).
545
Student’s t-test was performed for similar concentrations between light/dark for each
546
individual group (** - p < 0.01, *** - p < 0.001). B and C, Kinetics of ion leakage. Leaf
547
discs were treated with different concentrations of RB (B) and AO (C) for 0, 24, 48, 72 h
548
in the light. D and E, Histidine protection assay. Arabidopsis leaf discs were incubated
549
with 100 μM RB (D) or AO (E) in the presence or absence of 10 mM histidine (His), and
550
measured at 0, 2, 6, 24, 48 h. Means and SE are presented. The data was analyzed using
551
repeated measures two-way ANOVA (*** - p < 0.001).
552
553
Figure 3. Confocal microscopy of Arabidopsis seedlings treated with photosensitizers
554
AO and RB in the light (L) and the dark (D). Seedlings (5 day old, with tonoplast marker)
555
were treated with 100 μM of AO (A) or RB (B) and incubated for 72 h in the light or dark
556
and mounted on a microscope slide for observation by confocal microscopy. Scale bar is
557
20 μm.
558
559
Figure 4. Immunoblots of photosensitizer treated samples. Arabidopsis leaf discs from
560
AtSerpin1 overexpression (HA) lines were treated with 100 μM AO or RB or mock
20
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
21
561
treated (None) and incubated in the light for 0, 24, 48, 72 h. Extracts were treated with 50
562
μM E64 and fractionated on non-reducing SDS-PAGE and developed with α-AtSerpin1
563
antibody. The bottom images show the corresponding Coomassie staining.
564
565
Figure 5. Ion leakage assay of various Arabidopsis mutant lines treated with RB and AO.
566
Arabidopsis leaf discs were treated with 100 μM of RB (A) or AO (B) and incubated in
567
the light for 0, 24, 48, 72 h. Lines used were wild type (WT); AtSerpin1 overexpression
568
line (HA); AtSerpin1 insertion mutant (AtSerpin1-ko) and RD21 insertion mutant
569
(RD21-ko). The experiment was repeated three times, with means and SE presented. Data
570
was analyzed by repeated-measures two-way ANOVA. Significance established by
571
pairwise comparisons to the WT line using a post hoc Dunnett test (* - p < 0.05, *** - p <
572
0.001).
573
574
Figure 6. Ion leakage and singlet oxygen production during RB, AO and acute water
575
treatments. A, Ion leakage assay. Leaf discs from 10 week old WT (Col-0) plants were
576
subjected to various dehydration treatments for 0, 1, 2, 3 h and then rehydrated and
577
incubated under continuous light (30 μE). Readings were taken at 0, 0.5, 2, 6, 24, 48, 72
578
h. B, Confocal imaging of Arabidopsis seedlings subjected to dehydration. Seedlings with
579
the tonoplast marker (5 day old) were subjected to dehydration for 0, 15, 30 min, and
580
incubated with 100 μM SOSG in the dark, and mounted on a microscope slide for
581
observation by confocal microscopy as described in the Materials and Methods. Scale bar
582
is 20 μm. C, Ion leakage assay. Leaf discs were pre-incubated with 10 mM histidine,
583
subjected to dehydration treatment (0, 3 h) and incubated under continuous light (30 μE).
584
Readings were taken at 0, 2, 6, 24 h. The experiment was repeated twice, with means and
585
SE presented. The data was analyzed using repeated measures two-way ANOVA (** - p
586
< 0.01). D, EPR of RB, AO and dehydration-treated seedlings. WT seedlings were
587
incubated with 0.4 M 4-hydroxy-TEMP containing 0, 100, 200 and 400 μM RB or AO in
588
the dark for 30 min, washed and incubated in the light (30 μE) for 2 h. For dehydration,
589
WT Arabidopsis seedlings were incubated with 0.4 M 4-hydroxy-TEMP for 30 min,
21
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22
590
washed and then placed on dry Whatman paper for 0, 30, 60, 90 min in the dark.
591
Extraction was performed as described in the Materials and Methods before analysis by
592
EPR spectroscopy. The means and SE of 3 replicates is shown. Analyses by 2-way
593
ANOVA, followed by a post hoc Tukey test, or a Student’s t-test was performed against
594
their respective controls for significance (* - p < 0.05, ** - p < 0.01).
595
596
Figure 7. TBARS assay of RB, AO and dehydration treated Arabidopsis. WT
597
Arabidopsis seedlings were incubated with 100 μM of RB or AO for 1 h in the dark, then
598
washed and incubated in the light (30 μE) or dark for 0, 24 h. Dehydration treatment was
599
performed on WT Arabidopsis seedlings on dry Whatman paper or in water in the dark
600
for 0, 90 min. The TBARS assay was performed as described in the Materials and
601
Methods. The means and SE of 3 replicates is shown. Student’s t-test of the different
602
concentrations of RB or AO was performed against their respective controls for
603
significance (*- p < 0.05, ** - p < 0.01).
604
605
Figure 8. Arabidopsis mutant lines treated by dehydration stress. A, Immunoblot of
606
dehydration-treated plant AtSerpin1-HA. Leaf discs were treated on dry blotting paper
607
for 0 - 3 h as indicated, replaced in ddH2O for 1 h before harvesting. Extracts were
608
treated with 50 μM E64 and fractionated on non-reducing SDS-PAGE and developed
609
with α-AtSerpin1 antibody. The bottom image shows the corresponding Coomassie
610
staining. B, Ion leakage assay. Leaf discs from mutant plant lines as in Figure 5 were
611
subjected to 3 h dehydration treatment. Ion conductance readings were taken at 0, 24 h.
612
The experiment was repeated two times with means and SE presented. Student’s t-test of
613
each line against the WT line was performed for significance (* - p < 0.05).
614
615
Figure 9. Quantitative RT-PCR using 1O2 and O2-/H2O2 sensitive genes under various
616
treatments. A, Seedlings were incubated with 0, 0.1, 10 and 100 μM of RB, AO, MV in
617
the dark for 1 h, washed and then incubated in the light (30 μE) for 2 h. B, seedlings were
22
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
23
618
subject to 0, 30, 60, 90 min of dehydration in the dark. In MV treatment, numbers in
619
column represent off the scale values. The means and SE of 3 replicates is shown.
620
Student’s t-test was performed against their respective controls for significance (*- p <
621
0.05, ** - p < 0.01, ** - p < 0.001).
622
623
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24
624
625
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
A Tonoplast
RB/AO
Merge
B
(+)1M Mannitol
RB
AO
Figure 1. Confocal microscopy of Arabidopsis seedlings showing localization of
the photosensitizers RB and AO. A, Recombinant tonoplast marker line (5 day
old, CFP) treated with 100 µM RB (Top) and AO (Bottom) respectively. B,
Seedlings (5 day old, CFP) treated with 100 µM RB (Top) and AO (Bottom)
respectively, and mounted on a microscope slide with 1M mannitol to allow for
cell plasmolysis. Seedlings were observed by confocal microscopy as described
in the Material and Methods. Scale bar 20 µm.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
48 h
20
L
L
300 µM
0 µM
300 µM
0 µM
300 µM
D
D
0 µM
1 µM
10 µM
50 µM
100 µM
200 µM
300 µM
40
30
20
10
0
40
*
20
0
20
0 2 6 24 48 72
Time (h)
Time (h)
30
***
40
AO
+ His
60
0 µM
1 µM
10 µM
100 µM
300 µM
600 µM
1000 µM
60
0
E
- His
AO
0 2 6 24 48 72
D
RB
80
Ion leakage (%)
0 µM
300 µM
72 h
80
Ion leakage (%)
40
24 h
Ion leakage (%)
***
C
RB
50
**
60
0
B
RB
Ion leakage (%)
AO
0 µM
Ion leakage (%)
A
- His
+ His
20
***
10
0
2
6
24
Time (h)
48
2
6
24
48
Time (h)
Figure 2. Ion leakage in Arabidopsis plants showing effects of photosensitizers AO and RB in
the light and dark. A, Ion leakage assay. Leaf discs were treated with 300 µM AO and RB
respectively and incubated for 0, 24, 48, 72 hrs in the light (L) or dark (D). Student’s t-test was
performed for similar concentrations between light/dark for each individual group (** - p < 0.01,
*** - p < 0.001). B and C, Kinetics of ion leakage. Leaf discs were treated with different
concentrations of RB (B) and AO (C) for 0, 24, 48, 72 h in the light. D and E, Histidine
protection assay. Arabidopsis leaf discs were incubated with 100 µM RB (D) or AO (E) in the
presence or absence of 10 mM histidine (His), and measured at 0, 2, 6, 24, 48 h. Means and
SE are presented. The data was analyzed using repeated measures two-way ANOVA (*** - p
< 0.001).
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
A
Tonoplast
AO
Merge
Tonoplast
RB
Merge
L
D
B
L
D
Figure 3. Confocal microscopy of
Arabidopsis seedlings treated with
photosensitizers AO and RB in the light
(L) and the dark (D). Seedlings (5 day
old, with tonoplast marker) were treated
with 100 µM of AO (A) or RB (B) and
incubated for 72 h in the light or dark and
mounted on a microscope slide for
observation by confocal microscopy.
Scale bar is 20 µm.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
None
24 48 72
0
24
48 72
0 24 48 72
α-AtSerpin1
Light
0
RB
Complex (67 kD)
AtSerpin-HA(45 kD)
Coomassie
Time (h)
AO
Rubisco large subunit
(55 kD)
Spent serpin (37 kD)
Figure 4. Immunoblots of photosensitizer treated samples.
Arabidopsis leaf discs from AtSerpin1 overexpression (HA) lines were
treated with 100 µM AO or RB or mock treated (None) and incubated
in the light for 0, 24, 48, 72 h. Extracts were treated with 50 µM E64
and fractionated on non-reducing SDS-PAGE and developed with αAtSerpin1 antibody. The bottom images show the corresponding
Coomassie staining.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
A
Ion leakage (%)
40
30
WT
20
HA
10
AtSerpin1-ko
RD21-ko
0
0
24 48 72
Time (h)
B
Ion leakage (%)
40
30
*
20
***
10
WT
HA
AtSerpin1-ko
RD21-ko
0
0
24 48 72
Time (h)
Figure 5. Ion leakage assay of various
Arabidopsis mutant lines treated with RB
and AO. Arabidopsis leaf discs were
treated with 100 µM of RB (A) or AO (B)
and incubated in the light for 0, 24, 48,
72 h. Lines used were wild type (WT);
AtSerpin1 overexpression line (HA);
AtSerpin1 insertion mutant (AtSerpin1ko) and RD21 insertion mutant (RD21ko). The experiment was repeated three
times, with means and SE presented.
Data was analyzed by repeatedmeasures two-way ANOVA. Significance
established by pairwise comparisons to
the WT line using a post hoc Dunnett
test (* - p < 0.05, *** - p < 0.001).
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
A
B
Tonoplast
SOSG
Merge
Ion leakage (%)
80
0.5 h
60
0 min
2h
6h
40
24 h
20
15 min
48 h
72 h
0
30 min
0h 1h 2h 3h
80
60
**
0h
40
0.5 h
20
2h
0
6h
0h 3h 0h 3h
(-) His (+) His
24 h
80
60
*
**
* *
40
20
0
0 µM
100 µM
200 µM
400 µM
0 µM
100 µM
200 µM
400 µM
0 min
30 min
60 min
90 min
D
4-OH-TEMPO
concentration (nmol
g-1 FW)
Ion leakage (%)
C
RB
AO
Dry
Figure 6. Ion leakage and singlet oxygen production during RB, AO and acute water treatments. A, Ion
leakage assay. Leaf discs from 10 week old WT (Col-0) plants were subjected to various dehydration
treatments for 0, 1, 2, 3 h and then rehydrated and incubated under continuous light (30 µE). Readings were
taken at 0, 0.5, 2, 6, 24, 48, 72 h. B, Confocal imaging of Arabidopsis seedlings subjected to dehydration.
Seedlings with the tonoplast marker (5 day old) were subjected to dehydration for 0, 15, 30 min, and
incubated with 100 µM SOSG in the dark, and mounted on a microscope slide for observation by confocal
microscopy as described in the Materials and Methods. Scale bar is 20 µm. C, Ion leakage assay. Leaf discs
were pre-incubated with 10 mM histidine, subjected to dehydration treatment (0, 3 h) and incubated under
continuous light (30 µE). Readings were taken at 0, 2, 6, 24 h. The experiment was repeated twice, with
means and SE presented. The data was analyzed using repeated measures two-way ANOVA (** - p < 0.01).
D, EPR of RB, AO and dehydration-treated seedlings. WT seedlings were incubated with 0.4 M 4-hydroxyTEMP containing 0, 100, 200 and 400 µM RB or AO in the dark for 30 min, washed and incubated in the light
(30 µE) for 2 h. For dehydration, WT Arabidopsis seedlings were incubated with 0.4 M 4-hydroxy-TEMP for
30 min, washed and then placed on dry Whatman paper for 0, 30, 60, 90 min in the dark. Extraction was
performed as described in the Materials and Methods before analysis by EPR spectroscopy. The means and
SE of 3 replicates is shown. Analyses by 2-way ANOVA, followed by a post hoc Tukey test, or a Student’s ttest was performed against their respective controls for significance (* - p < 0.05, ** - p < 0.01).
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30
***
20
10
0
0h
24 h
AO Light
AO Dark
30
20
*
10
TBARS equivalents
(µmol g-1 FW)
RB Dark
TBARS equivalents
(µmol g-1 FW)
TBARS equivalents
(µmol g-1 FW)
RB Light
0
Wet
Dry
30
**
20
10
0
0h
24 h
0 min 90 min
Figure 7. TBARS assay of RB, AO and dehydration treated Arabidopsis. WT Arabidopsis
seedlings were incubated with 100 µM of RB or AO for 1 h in the dark, then washed and
incubated in the light (30 µE) or dark for 0, 24 h. Dehydration treatment was performed on WT
Arabidopsis seedlings on dry Whatman paper or in water in the dark for 0, 90 min. The TBARS
assay was performed as described in the Materials and Methods. The means and SE of 3
replicates is shown. Student’s t-test of the different concentrations of RB or AO was performed
against their respective controls for significance (*- p < 0.05, ** - p < 0.01).
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A
Dehydration
2
3
α-AtSerpin1
1
Complex (67 kD)
AtSerpin-HA (45 kD)
Spent serpin (37 kD)
Coomassie
Time (h) 0
Rubisco large
subunit (55 kD)
Ion leakage (%)
B
100
80
60
40
20
0
Dehydration 0 h
Dehydration 3 h
*
Figure 8. Arabidopsis mutant lines treated by dehydration stress. A, Immunoblot of
dehydration-treated plant AtSerpin1-HA. Leaf discs were treated on dry blotting paper
for 0 - 3 h as indicated, replaced in ddH2O for 1 h before harvesting. Extracts were
treated with 50 µM E64 and fractionated on non-reducing SDS-PAGE and developed
with α-AtSerpin1 antibody. The bottom image shows the corresponding Coomassie
staining. B, Ion leakage assay. Leaf discs from mutant plant lines as in Figure 5 were
subjected to 3 h dehydration treatment. Ion conductance readings were taken at 0, 24 h.
The experiment was repeated two times with means and SE presented. Student’s t-test
of each line against the WT line was performed for significance (* - p < 0.05).
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RB
AO
MV
Fold change Fold change Fold change
A
Control
20
10
0
20
***
***
***
*
10
0
*** *
1 µM
10 µM
***
***
***
***
***
*
**
*
*
**
*
At3g61190At5g64870At3g01830 At5g01600At4g21870At1g71030
*** ***
***
***
***
***
***
20
***
39 37
***
** ***
***
***
**
***
***
46
Fold change
Dry
Dry 0
20
10
21
O2- / H2O2
Dry 30
Dry 90
21
***
**
Dry 60
***
***
***
*
*** *
0
***
*** *
*
At3g61190 At5g64870 At3g01830 At5g01600 At4g21870 At1g71030
1O
2
B
100 µM
At3g61190 At5g64870 At3g01830 At5g01600 At4g21870 At1g71030
10
0
20
0.1 µM
***
** **
*** **
***
At3g61190 At5g64870 At3g01830 At5g01600 At4g21870 At1g71030
1O
2
O2- / H2O2
Figure 9. Quantitative RT-PCR using 1O2 and O2- / H2O2 sensitive genes under various
treatments. A, Seedlings were incubated with 0, 0.1, 10 and 100 µM of RB, AO, MV in
the dark for 1 h, washed and then incubated in the light (30 µE) for 2 h. B, seedlings
were subject to 0, 30, 60, 90 min of dehydration in the dark. In MV treatment, numbers
in column represent off the scale values. The means and SE of 3 replicates is shown.
Student’s t-test was performed against their respective controls for significance (*- p <
0.05, ** - p < 0.01, ** - p < 0.001).
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