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Abstract The double mutant ΔkatG/tpx of cyanobacterium Synechocystis sp. strain PCC 6803, defective in the anti-oxidative enzymes catalase (KatG) and thioredoxin peroxidase (Tpx), is unable to grow in the presence of exogenous H2O2. The ΔkatG/tpx mutant is shown to be extremely sensitive to very low concentrations of H2O2, especially when intensified with cold stress. Analysis of gene expression in both wild-type and ΔkatG/tpx mutant cells treated by combined cold/oxidative stress revealed that H2O2 participates in regulation of expression of cold-responsive genes, affecting either signal perception or transduction. The central role of a transmembrane stress-sensing histidine kinase Hik33 in the cold/oxidative signal transduction pathway is discussed. Introduction Abiotic stress signal transduction and response mechanisms are of special interest in fundamental research in cyanobacteria. Cyanobacteria are photosynthetic single-cell, filamentous or colony-forming prokaryotes that live in a variety of ecosystems, including freshwater and marine biomes. The survival of these organisms in environments with fluctuating physical and chemical properties such as temperature, light intensity or solute concentrations relies on immediate adaptive changes in cellular metabolism. Cyanobacterial reactions to stress are governed by alteration of gene expression; known cyanobacterial genomes include a number of genes that are transcribed and translated specifically in response to stress factors (Sinetova and Los 2016a). The single-cell cyanobacterium Synechocystis sp. strain PCC 6803 (hereafter Synechocystis), characterized by the optimal temperature for its growth at 30–36°C, is a model organism widely used in abiotic stress research. Change in transcription in response to cold stress has been studied and reviewed in detail (Los et al. 2008, Sinetova and Los 2016a). Usually, an incubation time period of 15–30 min at lowered temperatures is enough to detect changes in gene expression and confirm the cellular response to the stress (Mironov et al. 2012a). Cold stress-inducible genes are identified and can be divided into several major groups (Sinetova and Los 2016a): (i) fatty acid desaturases, necessary to introduce double bonds into fatty acids of phospholipids and reduce the viscosity of rigidified cell membranes; (ii) RNA-binding proteins and RNA helicases which support stress-induced transcription; (iii) ribosomal proteins which support translation at low temperatures; and (iv) proteases involved in repair of photodamaged PSII. In Synechocystis, the transcription of almost a half of all cold-inducible genes is regulated by a sensory histidine kinase, Hik33 (Suzuki et al. 2001). It is suggested that Hik33 may perceive changes in physical properties of the membrane lipid environment (such as microviscosity) via its transmembrane PAS (Per-Arnt-Sim) domains, which in response undergo conformational changes leading to autophosphorylation (Murata and Los 2006). Interestingly, in addition to membrane viscosity changes caused by lower temperatures, Hik33 phosphorylation is also sensitive to salt and hyperosmotic stress (Shumskaya et al. 2005), red light (Mironov et al. 2012b, Mironov et al. 2014) and oxidative stress (Kanesaki et al. 2007). Oxidative stress is characterized by the presence of reactive oxygen species (ROS) and accompanies various types of abiotic stresses including cold, strong light, high temperature or salinity (Schmitt et al. 2014). Most ROS have short half-lives which are hard to detect, with the exception of more stable hydrogen peroxide (H2O2). The effect of H2O2 on gene expression had been extensively studied in Synechocystis (Li et al. 2004, Kanesaki et al. 2007). Genome-wide transcriptomic analysis suggests that H2O2 acts as a universal molecule that triggers stress-induced gene expression (Sinetova and Los 2016b). Exogenously provided H2O2 induces expression of several groups of genes, and most of them also respond to a variety of stresses, for example: (i) heat stress protein-encoding genes (hspA, dnaJ, clpB1, dnaK2, ctpA, hypA1, htpG and sigB); (ii) genes induced by strong light and cold stress (hli); and (iii) genes involved in phycobilisome degradation and induced by multiple factors (nblA1 and nblA2). However, there is a group of genes specifically induced by H2O2. The products of these genes directly participate in ROS detoxification: catalase katG (sll1987), thioredoxin peroxidase tpx (sll0755), peroxiredoxin aphC (sll1621), glutaredoxins grxA (ssr2061), grxB (slr1562) and grxC (slr1846), as well as ferredoxin-plastoquinone reductase pgr5 (ssr2016). Genes induced by cold and/or oxidative stress are shown in Fig. 1. These genes, with a minor exception, respond to a signal from the Hik33 sensor (Suzuki et al. 2001, Kanesaki et al. 2007, Mironov et al. 2014). Also, while most of the genes respond to their specific stress, about 20% of them are induced by both cold and oxidative stress. This implies that their expression is triggered by a common factor present in both types of stress. We hypothesize that this common factor is H2O2 itself. Fig. 1 View largeDownload slide Genes induced by cold stress (22°C) and oxidative stress (0.25 mM of exogenous H2O2) in Synechocystis sp. strain PCC 6803. Expression of all genes is regulated by Hik33 (Suzuki et al. 2001, Kanesaki et al. 2007, Mironov et al. 2014) except those labeled with an asterisk. Genes in bold are those induced by both types of stress. Underlined genes are those used in qPCR analysis in this study. Fig. 1 View largeDownload slide Genes induced by cold stress (22°C) and oxidative stress (0.25 mM of exogenous H2O2) in Synechocystis sp. strain PCC 6803. Expression of all genes is regulated by Hik33 (Suzuki et al. 2001, Kanesaki et al. 2007, Mironov et al. 2014) except those labeled with an asterisk. Genes in bold are those induced by both types of stress. Underlined genes are those used in qPCR analysis in this study. Results Detection of expression of a specific gene induced by exogenous H2O2 presents a certain challenge. In Synechocystis, peroxide is immediately inactivated by anti-oxidative enzymes such as catalases or peroxidases; due to their action, it is quite difficult to detect any transcriptional changes specific to H2O2 addition. Thus, in order to test our hypothesis of H2O2 acting as a common factor altering gene expression during both cold and oxidative stress, we created a special double knock-out mutant of Synechocystis, ΔkatG/tpx. This mutant is deficient in a catalase gene katG (sll1987) and a thioredoxin peroxidase gene tpx (sll0755), the two genes specifically expressed in response to oxidation (Fig. 1). Due to the lack of detoxifying enzymes, the mutant was expected to have a delayed intracellular detoxification of H2O2 allowing a sufficient amount of time to detect specific gene expression. The ΔkatG/tpx mutant was subjected to a series of cold and oxidative stress treatments, and its physiological characteristics as well as the expression of genes were investigated; wild-type Synechocystis sp. strain PCC 6803 was used as a positive control. As a negative control, a mutant deficient in Hik33 (Δhik33, unable to receive and transduce major stress signals) was chosen. The choice was directed by the fact that both katG and tpx genes belong to the group of genes whose expression is governed by a signal from the Hik33 sensor (Suzuki et al. 2001, Kanesaki et al. 2007). H2O2 affects the growth of the ΔkatG/tpx mutant strain First, we evaluated the growth of the double mutant under standard and oxidative stress conditions. ΔkatG/tpx, as well as wild-type and Δhik33 cultures, were grown under the same standard conditions, and later subjected to oxidative stress as described in the Materials and Methods (Fig. 2). Under optimal conditions, the doubling time for all strains was similar: 10 ± 0.5 h for ΔkatG/tpx, 11 ± 0.8 h for Δhik33 and 9 ± 1.0 h for the wild type. Thus, under standard conditions, the double mutation did not cause any issues for the organism. Fig. 2 View largeDownload slide The effect of exogenous H2O2 on Synechocystis growth at 32°C. Point 0: H2O2 was added to a final concentration of 0.25 mM. Optical density measured at 750 nm reflects cell growth. Insert: ratio of absorbance at 680/750 nm, characteristic of the relative content of Chl in cells. The results are combined from four independent experiments. Fig. 2 View largeDownload slide The effect of exogenous H2O2 on Synechocystis growth at 32°C. Point 0: H2O2 was added to a final concentration of 0.25 mM. Optical density measured at 750 nm reflects cell growth. Insert: ratio of absorbance at 680/750 nm, characteristic of the relative content of Chl in cells. The results are combined from four independent experiments. However, after adding H2O2, ΔkatG/tpx mutant cells slowed in growth, remained alive for several hours and completely stopped growing by 8 h, in contrast to the wild-type and Δhik33 strains which continued to grow and gain biomass. The inability of the ΔkatG/tpx mutant to manage oxidative stress was also confirmed by the absorption spectrum of photosynthetic pigments taken for the strain throughout the experiment. The ratio of absorbance at 680/750 nm, characteristic for Chl content, showed a significant decrease indicative of Chl degradation (Fig. 2, insert). Indeed, the double mutant cell culture appeared blue, symptomatic of cell death. After 24 h of experimental treatment, the ΔkatG/tpx culture became colorless, pointing to its expected hypersensitivity to H2O2. Notably, the sensitivity to oxidative stress could be additionally exaggerated by the fact that BG-11 culture medium contains ferric iron. Addition of H2O2 into the medium supposedly initiates the Fenton reaction that produces hydroxyl radicals (Gutteridge 1991). While the wild-type strain and Δhik33 mutant were safeguarded by catalase and peroxidase activities, the double mutant had no protection against peroxide and hydroxyl radicals. The ΔkatG/tpx mutant strain fails to neutralize H2O2 In addition to the assessment of growth under oxidative stress, the rate of exogenous peroxide neutralization in cyanobacterial strains was tested using a peroxide assay. First, we assessed a normal, ‘steady-state’ amount of H2O2 in intact cells of the three strains under standard conditions. The steady-state concentration represents peroxide amounts accumulated in cells during conventional generation and destruction of ROS. It is known that autoclaved BG-11 growth medium contains trace amounts of H2O2 (20.3 ± 0.5 µM in our settings); thus, it was important to evaluate the ability of all three strains to retain minimal H2O2 when not exposed to any experimental stress. In the wild-type cells, the steady-state concentration was measured as 37 ± 3 µM; for Δhik33 it was 71 ± 6 µM and for ΔkatG/tpx cells it was 225 ± 15 µM. We have also attempted to remove any exogenous peroxide by washing the cells with water before measurements, which resulted in the following concentrations of H2O2: 42 ± 4 µM in the wild type, 89 ± 10 µM in the Δhik33 mutant and 45 ± 3 µM in the ΔkatG/tpx culture. Interestingly, while in the wild type the concentration of peroxide did not change significantly, suggesting a well-functioning metabolic system of ROS maintenance, and in the double mutant it expectedly and significantly decreased, the amount of peroxide in Δhik33 was twice as high as that in the wild type and did not change after washing. The high concentration of peroxide and the lack of change in its concentration in the Δhik33 mutant deficient in the major stress receptor indicates the presence of a mechanism that generates large amounts of intracellular H2O2 in Δhik33 cells. Next, the ability of strains to neutralize large amounts of exogenous H2O2 was tested by the peroxide assay. Peroxide was added to the washed cells to exclude possible errors introduced by residual ROS from the growth media, and its concentration in the cultures was measured at short (seconds to minutes) intervals (Fig. 3). As expected, in the wild-type strain, the concentration of H2O2 rapidly decreased almost to the base level of 35 ± 5 µM, agreeing with previously published data (Maeda et al. 2005). The peroxide half-life t1/2 for the wild type was measured as 21 ± 2 s, and the rate of peroxide degradation Vdeg/OD750 was estimated as 453 ± 14 µM min−1 (Table 1). In contrast, the Δhik33 mutant was characterized by a more prolonged peroxide inactivation caused by a lower degradation rate of Vdeg/OD750 = 173 ± 16 µM min−1 (t1/2 = 44 ± 3 s) and a higher concentration of H2O2 at the end of the experiment (115 ± 11 µM). Such results were anticipated since this mutant lacks the stress sensor and is unable to induce expression of genes for major anti-oxidative enzymes. Table 1 Degradation of H2O2 by cyanobacterial strains measured by the peroxide assay Parameters Strains Wild type Δhik33 ΔkatG/tpx C0 (µM) 35 ± 5 115 ± 11 44 ± 4 t1/2 21 ± 2 s 44 ± 3 s 8 ± 1 min Vdeg/OD750 (µM min−1) 453 ± 14 173 ± 16 6 ± 1 Parameters Strains Wild type Δhik33 ΔkatG/tpx C0 (µM) 35 ± 5 115 ± 11 44 ± 4 t1/2 21 ± 2 s 44 ± 3 s 8 ± 1 min Vdeg/OD750 (µM min−1) 453 ± 14 173 ± 16 6 ± 1 C0, base peroxide concentration, t1/2 = te×ln2 (peroxide half-life, te refers to e-folding time parameter), Vdeg, C×te−1 (rate of peroxide oxidation, C refers to the peroxide concentration). Note: C0 is measured in cells washed with water and does not reflect the natural concentration of H2O2 in an intact growth culture. Table 1 Degradation of H2O2 by cyanobacterial strains measured by the peroxide assay Parameters Strains Wild type Δhik33 ΔkatG/tpx C0 (µM) 35 ± 5 115 ± 11 44 ± 4 t1/2 21 ± 2 s 44 ± 3 s 8 ± 1 min Vdeg/OD750 (µM min−1) 453 ± 14 173 ± 16 6 ± 1 Parameters Strains Wild type Δhik33 ΔkatG/tpx C0 (µM) 35 ± 5 115 ± 11 44 ± 4 t1/2 21 ± 2 s 44 ± 3 s 8 ± 1 min Vdeg/OD750 (µM min−1) 453 ± 14 173 ± 16 6 ± 1 C0, base peroxide concentration, t1/2 = te×ln2 (peroxide half-life, te refers to e-folding time parameter), Vdeg, C×te−1 (rate of peroxide oxidation, C refers to the peroxide concentration). Note: C0 is measured in cells washed with water and does not reflect the natural concentration of H2O2 in an intact growth culture. Fig. 3 View largeDownload slide Kinetics of degradation of exogenous H2O2 by Synechocystis strains. Time 0: addition of H2O2 to washed cells to a final concentration of 0.25 mM. Aliquots of treated cells were taken at time intervals of 4 s, 8 s, 15 s, 30 s, 1 min, 2 min, 4 min and 8 min, and the concentration of H2O2 was measured. The experiment was performed in three biological replicates. Overall, four independent experiments were conducted. Fig. 3 View largeDownload slide Kinetics of degradation of exogenous H2O2 by Synechocystis strains. Time 0: addition of H2O2 to washed cells to a final concentration of 0.25 mM. Aliquots of treated cells were taken at time intervals of 4 s, 8 s, 15 s, 30 s, 1 min, 2 min, 4 min and 8 min, and the concentration of H2O2 was measured. The experiment was performed in three biological replicates. Overall, four independent experiments were conducted. The ΔkatG/tpx mutant demonstrated the slowest degradation of peroxide. For this mutant, the average concentration of peroxide remained the highest of the three strains during the 8 min of the experiment, implying that the mutant was unable to detoxify peroxide effectively (Vdeg/OD750 = 6 ± 1 µM min−1) (Table 1). The results were concordant with the lack of two major anti-oxidative enzymes, catalase and thioredoxin peroxidase. Light intensifies H2O2 accumulation in Δhik33 and ΔkatG/tpx under cold stress Light is another factor that naturally increases the amount of ROS in photosynthetic cells (Mironov et al. 2012b). We hypothesized that in the dark the double mutant would accumulate and retain much less but a still significant amount of H2O2, thus surviving cold stress or combined cold and oxidative stress better. In order to test our hypothesis, we subjected the wild type, Δhik33 and ΔkatG/tpx strains to cold stress and compared the amount of steady-state H2O2 during light (120 µmol photons m−2 s−1) or dark incubation (Fig. 4). Indeed, we found that for both mutant strains the concentration of H2O2 increased when cold stress was provided in the presence of light. However, the amount of H2O2 was still acceptable in both cases during 32 min of treatment, so for comparability we decided not to alter light conditions for our stress treatments. Fig. 4 View largeDownload slide Changes in the steady-state H2O2 concentration in Synechocystis strains under cold stress in the presence or absence of light (120 μmol quanta m−2 s−1). Wild-type, Δhik33 and ΔkatG/tpx cultures of Synechocystis were initially grown at 32°C and then shifted to 22°C in the light (open symbols) or in the dark (filled symbols) for 32 min. H2O2 concentrations were measured for unwashed cells after 2, 4, 8, 16 and 32 min of cold treatment. The results are combined from three independent experiments. Fig. 4 View largeDownload slide Changes in the steady-state H2O2 concentration in Synechocystis strains under cold stress in the presence or absence of light (120 μmol quanta m−2 s−1). Wild-type, Δhik33 and ΔkatG/tpx cultures of Synechocystis were initially grown at 32°C and then shifted to 22°C in the light (open symbols) or in the dark (filled symbols) for 32 min. H2O2 concentrations were measured for unwashed cells after 2, 4, 8, 16 and 32 min of cold treatment. The results are combined from three independent experiments. H2O2 regulates the expression of genes induced by both cold and oxidative stress Once we confirmed that the ΔkatG/tpx mutant can survive a short period of oxidative stress with continuous illumination while retaining significant amounts of H2O2, we proceeded with the assessment of transcription induced by cold and peroxide simultaneously. All three Synechocystis cultures were initially grown under optimal conditions (32°C), and then diluted and separated into three experimental culture tubes (Fig. 5A). A small aliquot of cells was taken from all three tubes, mixed and used later as the control (untreated sample at 32°C). Once control samples were removed, the remaining tubes were immediately subjected to the following treatments: (i) to the first tube, H2O2 was added to a final concentration of 0.25 mM; (ii) in the second tube, the temperature was lowered to 22°C; and (iii) the third tube was subjected to a combined treatment of H2O2 (0.25 mM) and low temperature (22°C). Fig. 5 View largeDownload slide Analysis of stress-inducible gene transcription in Synechocystis strains. (A) Scheme of the experiment. Each strain [wild type (WT), Δhik33 and ΔkatG/tpx] was initially grown at 32°C to OD750 = 2 before being split into three experimental tubes. The control samples were collected from all three tubes and mixed; immediatley after that the experimental tubes were subjected to treatments. Peroxide was added to the first tube to a final concentration of 0.25 mM; the second tube was moved to 22°C; the third tube was subjected to a combined stress of peroxide and low temperature. After 30 min, cells were fixed and used for isolation of total RNA. (B) Analysis of transcription of some stress-inducible genes that are listed in Fig. 1. Bars represent relative expression of the genes; amounts of transcripts were assessed by qRT–PCR, normalized to those of rnpB and expressed as a percentage of the maximum value of gene expression in all samples. +, with addition of H2O2; –, without addition of H2O2. Fig. 5 View largeDownload slide Analysis of stress-inducible gene transcription in Synechocystis strains. (A) Scheme of the experiment. Each strain [wild type (WT), Δhik33 and ΔkatG/tpx] was initially grown at 32°C to OD750 = 2 before being split into three experimental tubes. The control samples were collected from all three tubes and mixed; immediatley after that the experimental tubes were subjected to treatments. Peroxide was added to the first tube to a final concentration of 0.25 mM; the second tube was moved to 22°C; the third tube was subjected to a combined stress of peroxide and low temperature. After 30 min, cells were fixed and used for isolation of total RNA. (B) Analysis of transcription of some stress-inducible genes that are listed in Fig. 1. Bars represent relative expression of the genes; amounts of transcripts were assessed by qRT–PCR, normalized to those of rnpB and expressed as a percentage of the maximum value of gene expression in all samples. +, with addition of H2O2; –, without addition of H2O2. All treatments continued for exactly 30 min, after which cells were fixed and used for the isolation of total RNA (Fig. 5A). Relative amounts of specific mRNAs for the genes normally responsive to both cold and oxidative stress (desB, ndhD2, hliB and pgr5), as well as an H2O2-inducible sodB and a Hik33-independent gene rbpA (Fig. 1) were estimated by quantitative reverse transcription–PCR (qRT–PCR) (Fig. 5B). As expected, the transcription of desB (encoding omega-3 fatty acid desaturase) was induced by cold stress in both the wild type and the double mutant, while no significant desB induction was detected in the Δhik33 mutant, in agreement with the lack of the stress signal receptor. For the cold stress, induction factors calculated as the ratio between levels of transcripts in stressed and control cells were 5.8 ± 0.2 for the wild type and 11 ± 1 for the ΔkatG/tpx double mutant. Interestingly, while in the wild type the expression of desB was not drastically affected by the presence of H2O2, in the ΔkatG/tpx mutant it was significantly inhibited by the exogenous H2O2. This effect was present at both temperatures: at 32°C the induction factor of desB was 0.06 ± 0.03 (a 17-fold decrease compared with the peroxide-free environment); at 22°C it was 0.2 ± 0.03 (a drastic 55-fold decrease). Thus, in the double mutant under cold stress H2O2 operated as a negative regulator for desB induction. NdhD2 was regulated similarly to desB, except in the double mutant there was a small induction under the combined peroxide and cold treatments. Transcription of hliB was sensitive to both cold stress and H2O2. In the wild type, cold stress caused about a 20-fold accumulation of the gene transcript, while the combined treatment with cold and H2O2 led to a significant 120-fold increase. In the ΔkatG/tpx mutant, H2O2 alone caused a 173-fold accumulation, cold treatment resulted in 22-fold accumulation and the combined stress caused a drastic 380-fold increase. Thus, H2O2 positively regulated the expression of hliB in a Hik33-dependent manner. Transcription of pgr5 was regulated similarly to hliB. Cold induction of cold-inducible rbpA was confirmed to be Hik33 independent, and all three strains demonstrated the expression of this gene. However, the ΔkatG/tpx mutant was more sensitive to cold in the presence of H2O2. The sodB gene encoding a superoxide dismutase was strongly induced in the wild-type cells only under combined action of cold and exogenous H2O2. In ΔkatG/tpx cells, the induction of sodB by H2O2 was detected even at a normal growth temperature. This gene is also regulated by Hik33, so its expression was absent in the Δhik33 mutant, as expected. Discussion Role of Tpx and KatG in detoxification during oxidative stress Previously, it was shown that KatG catalase and Tpx peroxidase maintain intracellular concentrations of H2O2 at low, non-toxic levels. For example, a simultaneous knock-out of both katG and tpx genes in Synechocystis prevents repair of photodamaged PSII in the presence of ROS (Nishiyama et al. 2001), suggesting anti-oxidant properties of these enzymes. However, not much data was available regarding their specific functions. A single knock-out of either katG (Tichy and Vermaas 1999) or tpx (Yamamoto et al. 1999) does not affect the growth of Synechocystis under standard conditions, suggesting that neither of the genes is essential for growth under non-oxidative conditions; in ΔkatG, however, the rate of H2O2 decomposition decreases 30-fold. Furthermore, the Δtpx mutant was also unable to quench peroxide-dependent Chl fluorescence, supporting the hypothesis that Tpx is the only peroxidase in Synechocystis that scavenges H2O2 and alkyl hydroperoxides (such as tertiary butyl hydroperoxide) with thioredoxin as an electron donor. In the present study under standard conditions, the ΔkatG/tpx mutant did not demonstrate any difference in growth in comparison with wild-type Synechocysits, supporting the hypothesis that neither of the genes is essential. The importance of both genes became evident once we demonstrated that cells of the double mutant cannot metabolize the excess exogenous H2O2 (Table 1). We have confirmed that both KatG and Tpx are involved in H2O2 detoxification. The inability of the katG/tpx mutant to detoxify H2O2 rapidly makes it an excellent model to study the effects of exogenous H2O2 on gene expression (especially in the presence of cold stress). It should be noted that exogenous H2O2 can be transported through a membrane passively down the concentration gradient or actively via aquaporins, and should be distinguished from an intracellular H2O2 which is a result of metabolic reactions. Migration of H2O2 molecules through cell membranes occurs in seconds (Bienert et al. 2006); in the present study, we did not discriminate between intracellular and extracellular peroxide species assuming that extra- and intracellular peroxide concentrations were in a steady-state equilibrium, since the measurements of peroxide concentration were conducted at intervals of seconds to minutes. Hik33 participates in sensing and regulation of intracellular H2O2 Under standard conditions, BG-11 medium always contains a residual amount of H2O2 generated during autoclaving. In order to account for that, some BG-11 recipes recommend addition of a reducing agent such as sodium thiosulfate while preparing a cyanobacterial culture (Zhang et al. 2004). Thus, we had to wash our cultures with deionized water before the peroxide assay to eliminate any input from the residual exogenous ROS. Surprisingly, washing of the Δhik33 mutant culture did not result in a significant decrease of the steady-state peroxide concentration. We hypothesized that Δhik33 cells could produce a significant amount of peroxide and expel it to the extracellular space. While this hypothesis needs thorough testing, we considered the possibility that the excessively produced H2O2 results from lowered activities of katG and tpx unable to receive a stress signal from missing Hik33, which is believed to be an H2O2 receptor (as shown by Kanesaki et al. 2007). According to the data on the transcription intensity of katG and tpx (Fig. 6), determined from our qRT–PCR data, Δhik33 cells have >2-fold the transcript levels of these genes compared with the wild type under standard conditions. Δhik33 thus apparently has sufficient katG and tpx transcripts to detoxify peroxide and should have had a reduced amount of steady-state peroxide after washing; the presence of peroxide in the cells even after the residual peroxide is washed out suggests an unknown mechanism of ROS generation in the Δhik33 mutant. Fig. 6 View largeDownload slide Relative expression of katG and tpx in the wild type and Δhik33 mutant of Synechocystis grown under standard conditions. Amounts of transcripts were assessed by qRT–PCR and normalized to those of rnpB. Fig. 6 View largeDownload slide Relative expression of katG and tpx in the wild type and Δhik33 mutant of Synechocystis grown under standard conditions. Amounts of transcripts were assessed by qRT–PCR and normalized to those of rnpB. H2O2 is a common regulator present in both cold and oxidative stress Here, we presented data showing that H2O2 is a factor regulating the cellular response to cold stress. Due to a rapid detoxification of peroxide in normal conditions, the minute effect of H2O2 on gene expression can be noticed only in a mutant lacking major anti-oxidative enzymes. We have shown that such a mutant has a delayed detoxification mechanism which gives plenty of time to detect changes in gene expression. We discovered that in ΔkatG/tpx cells, while the amount of desB or ndhD2 transcripts increased due to the cold stress, it significantly decreased when exposed to combined cold and oxidative stress. This suggests that H2O2 negatively regulates cold-dependent transcription of these genes. Interestingly, functions of both desB and ndhD2 are directly or indirectly related to the physical state of membranes. The desB gene encodes an omega-3 fatty acid desaturase, which lowers the melting temperature of phospholipids, affecting membrane fluidity. The connection between ndhD2 and the membrane state is more indirect: NdhD2 is a subunit of NADPH-quinone oxidoreductase, which acts as an acceptor of electrons from NADPH in both cyclic and respiratory electron transport chains (Battchikova et al. 2011). Hyperactive NdhD2 would decrease the amount of NADPH which is also required for fatty acid biosynthesis, and affect the redox state of a quinone pool which is linked to membrane fluidity (Maksimov et al. 2017). In contrast, the expression of hliB and pgr5 genes is significantly increased if exposed to combined cold and peroxide stress, implying that H2O2 is a positive regulator of cold-independent transcription. The products of both genes regulate the activity of the photosynthetic apparatus. HliB participates in a stress-induced recovery process of damaged components of PSII via binding to Chl during its assembly and repair (Promnares et al. 2006). Pgr5 is a protein that participates in cyclic electron flow during a shortage of reducing donors (Munekage et al. 2002, Yeremenko et al. 2005, Yamamoto et al. 2016). Both proteins attenuate the effects of photoinhibition caused by exposure of cells to strong light. Assuming the UV light present in strong light increases formation of ROS, the stimulation of hliB and pgr5 transcription by H2O2 is anticipated. The rbpA gene is a typical representative of a group of cold-induced, Hik33- and light-independent genes (Mironov et al. 2012b). We did not find any enhancement of its transcription by the combined stress action in the wild type; however, in the double mutant, both exogenous peroxide and cold stress led to the 2-fold induction of this gene. More work is needed to position this phenomenon in the framework of signaling systems of Synechocystis. The sodB gene encodes a superoxide dismutase (FeSOD) that catalyzes the conversion of superoxide into peroxide and may generate endogenous H2O2. SodB is a Hik33-dependent gene and is particularly induced by the combined stress in both the wild type and the ΔkatG/tpx mutant (Fig. 5). While our results on cold stress transcription are consistent with the results obtained earlier (Suzuki et al. 2001, Mironov et al. 2012b, Mironov et al. 2014), we have discovered a unique connection between peroxide and cold stresses. Our results suggest that H2O2 is a common factor present in both types of stress, regulating the expression of cold-inducible genes. In our study, the ΔkatG/tpx mutant was extremely sensitive to very small amounts of H2O2, altering gene expression correspondingly. We propose that H2O2 interferes with the cold regulation process during signal perception or transduction, either enhancing the effect of cold or, conversely, reducing it. Hik33 plays a central role in ROS-mediated signal detection for light, cold and oxidative stress Earlier we demonstrated that the induction of Hik33-mediated cold shock genes is regulated by light (Mironov et al. 2012b,, Mironov et al. 2014). Here, we show that there is an additional regulator: ROS (H2O2). Such a connection would be anticipated since in photosynthetic cells various ROS are formed under light as a result of regular activity of both PSII and PSI. We suggest that Hik33 operates in a light- and ROS-dependent manner, forming a regulatory physiological circuit in cyanobacterial cells, and the light plays an important role in ROS metabolism (Fig. 4). The mechanism of ROS production in photosynthesis differs between photosystems and organisms. In PSI, a univalent reduction of O2 by electrons from PSII generates superoxide radicals · O2−, which are converted to H2O2 and O2 by superoxide dismutase, with H2O2 later reduced to water by catalases and peroxidases (Asada 2006). Generation of the superoxide anion as a primary product on the donor side of PSI (the Mehler reaction) is well described in chloroplasts of algae and higher plants (Badger et al. 2000). In cyanobacteria, however, the extent of the Mehler reaction is still questionable. Some reports show that at least in a model Synechocystis, the Mehler-like reaction does not produce ROS due to flavodiiron proteins that safely forward electrons released from water splitting by PSII directly to O2, reducing it to water in a water–water cycle (Helman et al. 2003, Allahverdiyeva et al. 2011, Allahverdiyeva et al. 2013). However, the genes for those flavodiiron proteins can be easily knocked out, and the phenotypes of the resultant mutants do not differ dramatically from those of the wild-type cells. In contrast, the knockout of the sodB gene for the FeSOD is either impossible under regular photoautotrophic conditions (Nefedova et al. 2003, Ke et al. 2014), or requires low light intensity and supplementation of the growth medium with sodium bicarbonate and catalase or bovine serum albumin (Nefedova et al. 2003). These observations point to the participation of the plant-type water–water cycle in ROS generation in Synechocystis. In PSII the mechanism of ROS generation is more similar between eukaryotes and prokaryotes. Singlet oxygen is produced by the energy input to O2 from photosensitized Chl. ROS is most probably formed by reducing species at the electron acceptor side and highly oxidizing components at the electron donor side (Pospíšil 2009). On the donor side of PSII, two-electron oxidation of H2O by the water-splitting manganese complex results in the formation of H2O2. On the acceptor side of PSII, ROS (superoxide anion radicals, H2O2 and hydroxyl radicals) are generated by a series of reactions starting with the reduction of molecular oxygen supported by plastoquinones (Lambreva et al. 2014, Khorobrykh et al. 2015). Our observation that a combined effect of cold stress and light increases the amount of H2O2 in cells is in agreement with a number of previous reports that demonstrate stress-induced ROS accumulation under light (reviewed in Dietz et al. 2016). Additionally, the light-dependent accumulation of ROS is demonstrated by the experiments in Chlamydomonas reinhardtii conducted by Michelet et al. (2013). Their research showed that under high light the amount of H2O2 transiently increases to induce stress-specific gene expression, while catalase activity decreases. It was concluded that it is the redox state of plastoquinones that is somehow sensed under strong light and used as a signal to decrease the catalase activity, allowing transient accumulation of H2O2 to induce subsequent signaling events (Michelet et al. 2013). In Synechocystis, a direct interaction between H2O2 and either Hik33 or transcription factors cannot be excluded. As known from the literature, the H2O2 redox signal per se is an oxidation of thiol groups of cysteine residues in a protein, followed by a change of the protein conformation. Oxidation of proteins by excessive H2O2 triggers a cascade of subsequent molecular events (Marinho et al. 2014). In contrast to multiple transcription factors which indeed may utilize this mechanism, Hik33 in Synechocystis does not contain a single cysteine residue. However, Hik33 contains two domains, PAS (Per-Arnt-Sim) and HAMP (histidine kinases, adenylyl cyclases, methyl-accepting chemotaxis proteins, phosphatases) (Khursigara et al. 2008), which may function as input and output modules for signal transduction in a redox-mediated manner. Since PAS domain-containing polypeptides are associated with light-dependent electron transport chains (Tayor and Zhulin 1999), it is possible that they respond to ROS (H2O2) signaling mediated by redox changes in photosynthetic quinones. Materials and Methods Cyanobacterial cultures and treatments Synechocystis sp. strain PCC 6803 substrain GT (glucose tolerant) was used as the wild type. Synechocystis culture was maintained using BG-11 agarized medium (Rippka et al. 1979) buffered with 20 mM HEPES-NaOH pH 7.5. For all experiments, culture growth was started at an initial OD750 = 0.2 and optimized at illumination with 100–120 μmol quanta m−2 s−1, final CO2 concentration in air of 1.5% and incubation temperature of 32°C in liquid BG11 medium with no antibiotics. Stress inductions were performed during the exponential growth at OD750 = 2. For a combined cold/dark treatment, all cultures were grown as described above, at OD750 = 2. The temperature was lowered to 22°C and cultures were split in half; one left in the light, and one moved to darkness. For a cold treatment, cultures were moved to 22°C. For the oxidative stress, the treatment consisted of adding of H2O2 to the cultures at a final concentration of 0.25 mM. Mutant construction The mutant strain deficient in hik33 (Δhik33) was produced previously (Suzuki et al. 2001, Mironov et al. 2014). A double ΔkatG/tpx knock-out mutant was constructed using target mutagenesis (Supplementary Fig. S1). Gene cassettes encoding resistance to the antibiotics spectinomycin (from pAM1303; Kulkarni and Golden 1997) and kanamycin (from pUC4-KIXX, Pharmacia) were inserted into sequences of the corresponding genes in the Synechocystis genome. First, genomic DNA was isolated from Synechocystis as described previously (Mironov and Los 2015). The nucleotide sequence of katG was amplified from genomic DNA using Taq polymerase (Thermofisher) and primers katG-F and katG-R (Supplementary Table S1). The resulting PCR fragment was cloned into the pTZ57R vector. Both the vector and spectinomycin cassette (SpR) were treated with NcoI, and then ligated so that the SpR was inserted in the middle of the coding sequence of katG. The resulting construct was used to transform Synechocystis, allowing recombination and insertion of the mutated gene into the genome. Successful transformants were selected using solid growth media containing spectinomycin. The presence of the insertion was confirmed by an increased size of the gene in genomic DNA detected by PCR. Similarly, the nucleotide sequence of the tpx gene was amplified with tpx-F and tpx-R primers (Supplementary Table S1) and cloned into pTZ57R. A kanamycin resistance cassette (KmR) was inserted into the coding sequence of tpx. The plasmid containing the tpx gene with KmR was used to transform the culture of the Synechocystis ΔkatG mutant described above. The resulting double mutant ΔkatG/tpx was able to grow in the presence of two antibiotics, kanamycin and spectinomycin. Complete substitution of the original genes in several copies of Synechocystis chromosomes with mutated genes was achieved through continuous passages and an increase in the antibiotic concentration. Complete substitution of the native genes with the mutated genes was confirmed by PCR (Supplementary Fig. S1). The katG/tpx-deficient mutant was maintained on solid BG11 medium supplemented with 25 μg ml−1 kanamycin and 30 μg ml−1 spectinomycin. RNA isolation and qRT–PCR Total RNA was isolated as described previously (Mironov and Los 2015). Primer selection for qRT–PCR was performed using the Geneious R8.1 program (https://www.geneious.com) (Supplementary Table S1). cDNA was synthesized using 1 μg of total RNA, a degenerate octameric primer (Evrogen Ltd.) and MMLV reverse transcriptase (Evrogen). The reaction was performed at 42°C for 1 h and stopped by a 10-fold dilution of deionized water. qRT–PCR was performed in a CFX96 Touch Real-Time PCR Detection System (Bio-Rad) using qPCRmix-HS SYBR pre-mix (Evrogen). The qRT–PCR protocol comprised pre-heating samples for 5 min at 95°C, followed by 45 cycles of a two-step amplification of 95°C (15 s) and 63°C (45 s). The fluorescence intensity of SYBR was taken at 63°C. At the end of the PCR, melting points of the PCR products were tested. The samples were incubated at various temperatures (65–90°C) for 5 s in 1°C steps. The first derivative of the obtained melting curves was used to evaluate the homogeneity of the fragments obtained during PCR. All reactions were characterized by the presence of exactly one peak with a maximum at >80°C, which corresponded to the amplification of the target products during PCR. Transcript levels of genes were normalized to those of rnpB by the ΔΔCt method. Amplification efficiency for each reaction was corrected using a calibration curve. For each gene, the experiments were performed in two biological and three analytical replicates. For each gene, expression values were presented as a percentage assuming the maximum level of induction in one experiment to be 100%. Measurement of H2O2 content H2O2 content was measured using a modification of the procedure of Graf and Penniston (1980). A 100 μl aliquot of cyanobacterial culture was mixed with 1 ml of a reaction solution prepared fresh before the experiment [1 ml of 50 mM HCl, 100 μl of 1 M KI, 100 μl of 1.0 mM ammonium molybdate in 0.5 M H2SO4 and 100 μl of 5% (w/v) starch]. The resulting 1.1 ml solution was thoroughly mixed and centrifuged for 1 min at 16,000×g. The supernatant was collected and incubated at 37°C for 20 min, then the absorbance of the solution was measured at 570 nm. H2O2 concentration was determined using a standard curve. The H2O2 standard curve was prepared by measuring the absorbance of 2-fold serial dilutions of 0.003% H2O2 (i.e. 882 μM, 441 μM, 221 μM, 110 μM, etc.) in the reaction solution. Peroxide assay Intracellular peroxidase activity was indirectly assessed using a peroxide assay. A 10 ml aliquot of culture was centrifuged at 3,000×g then washed by distilled water to remove residual H2O2. Pelleted cells were resuspended in fresh water at OD750, and a base peroxide concentration (C0) was measured. Once the base concentration was estimated, an additional amount of peroxide solution was added to a final concentration of 0.25 mM. Peroxide concentrations (C) were measured at designated intervals using the method described above. The results were plotted as concentration vs. time and the curve was identified in SciDAVis 1.D009 (http://scidavis.sourceforge.net/) as a first-order exponential decay. Peroxide half-life was calculated as t1/2 = te×ln2, where te is the e-folding time parameter. The rate of peroxide degradation was calculated as Vdeg = C×te−1 (µM min−1). Vdeg was normalized to OD750. For all calculations, R2 was >0.99. Supplementary Data Supplementary data are available at PCP online. Funding This work was supported by the Russian Science Foundation [grant No. 14-24-00020 to D.A.L.]. Disclosures The authors have no conflicts of interest to declare. 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Plant and Cell Physiology – Oxford University Press
Published: Mar 24, 2018
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