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Applied and Environmental Microbiology, November 2008, p. 6521-6527, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.01481-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Nitric Oxide as a Signaling Factor To Upregulate the Death-Specific Protein in a Marine Diatom, Skeletonema costatum, during Blockage of Electron Flow in Photosynthesis

Chih-Ching Chung,^1,2 Sheng-Ping L. Hwang,³ and Jeng Chang^2,4*

Center for Marine Bioscience and Biotechnology, National Taiwan Ocean University, Keelung 20224, Taiwan, Republic of China,¹ Institute of Marine Environmental Chemistry and Ecology, National Taiwan Ocean University, Keelung 20224, Taiwan, Republic of China,² Institute of Cellular and Organismic Biology, Academia Sinica, Nankang, Taipei 11529, Taiwan, Republic of China,³ Institute of Marine Biology, National Taiwan Ocean University, Keelung 20224, Taiwan, Republic of China⁴

Received 2 July 2008/ Accepted 26 August 2008

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To determine the physiological functions of a novel death-specific protein gene, Skeletonema costatum DSP-1 (ScDSP-1) in a marine diatom, Skeletonema costatum, the mRNA abundance of ScDSP-1 was measured in cultures subjected to light manipulation and treatments with various chemicals. When cells were transferred to a dim light intensity of 15 µmol m^–2 s^–1, ScDSP-1 mRNA levels showed a transient increase of 1 to 17.2 µmol (mol 18S rRNA)^–1 in 60 h. Furthermore, treatments with the photoinhibitors 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) and 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB) resulted in high ScDSP-1 mRNA levels, which reached 943 and 72 µmol (mol 18S rRNA)^–1, respectively. Treatment with the nitric oxide (NO) donor diethylamine nitric oxide also induced ScDSP-1 expression, and this inducible expression was inhibited by the NO scavenger hemoglobin. Additionally, the expression of ScDSP-1 mRNA elicited by DCMU and DBMIB was efficiently reduced when cultures were pretreated with the cell-penetrating NO scavenger 2-(4-carboxyphenyl)-4,5-dihydro-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide. In contrast, treatment with another photoinhibitor, paraquat, had no effect on ScDSP-1 expression. Our results indicated that NO is the crucial secondary messenger which signals the expression of ScDSP-1 when electron flow between photosystem II and photosystem I is blocked in S. costatum cells. In addition, the discovery of a similar gene, ScDSP-2, is briefly described.

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Diatoms are one of the most important groups of phytoplankton responsible for global primary productivity (13, 21, 23, 54). Determining the factors that regulate the succession of diatom populations has important implications in the study of energy flow and nutrient cycling in marine ecosystems. At the end of a diatom bloom, massive cell loss usually occurs. In addition to sedimentation and grazing by herbivores, programmed cell death (PCD) of stressed cells is also considered one of the major causes for the decline of algal blooms (8, 9, 56). Such a PCD process is triggered mainly by external stress factors, such as nutrient starvation or light deprivation (5, 50). In addition, virus infection (8, 25) and carbon dioxide limitation (56) are other factors resulting in PCD.

The process of PCD executed by a superfamily of cysteine aspartate-specific proteinases (caspases) is a conserved mechanism of cell suicide (36, 53). According to cell morphology and biochemical assays, autolysis in stressed phytoplankton is considered to be a process analogous to PCD which occurs in metazoans (6, 8, 24, 50). How the signal transmission of environmental stresses eventually elicits PCD in phytoplankton is an intriguing question. Among environmental factors, light availability is considered the most important factor for phytoplankton growth. Previous studies indicated that light deprivation triggered the PCD process in algal cells (5, 8, 50). For example, darkness resulted in PCD in a chlorophyte alga, Dunaliella tertiolecta, during which a decline in the photosynthesis efficiency induced a set of caspase-like proteases (50). On the other hand, the induction of reactive oxygen species (ROS) by high irradiance has been reported to trigger PCD in a marine cyanobacterium, Trichodesmium (6).

ROS and nitric oxide (NO) are now known to be important messengers of stress responses in plants (12, 26, 29). ROS is a by-product of photosynthesis, especially under conditions that cause the electron transport chain to go into overdrive (3, 42). The accumulation of ROS in chloroplasts initiated PCD in guard cells of leaves of the pea, Pisum sativum (48). In addition, depletion of dissolved carbon dioxide led to the formation of ROS and induced PCD-like autolysis at the end of a dinoflagellate bloom of Peridinium gatunense (56). On the other hand, treating a suspension cell culture of the orange plant, Citrus sinensi, with individual NO donors caused cessation of cell growth and subsequently induced the PCD process (49). Similarly, NO promoted soybean cell death by reactive oxygen intermediates when a bacterium, Pseudomonas syringae, infected the plant (14). In marine diatoms, such as Thalassiosira weissflogii and Phaeodactylum tricornutum, a group of diatom-derived aldehyde products has been demonstrated to induce calcium-dependent NO synthase-like activity, and this result implies that NO and calcium ions are secondary messengers participating in signal transduction in the PCD regulatory pathway (57). Moreover, blockage of photosynthetic electron transport activity either by darkness or by photosystem (PS) inhibitors, such as 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), is required for NO production in several species, including two green algae, Chlamydomonas reinhardtii and Scenedesmus obliquus, and a cyanobacterium, Anabaena doliolum (38, 47).

Recently, a novel gene, Skeletonema costatum DSP-1 (ScDSP-1) (formerly termed ScDSP) (11), encoding a death-specific protein (DSP) was obtained from a marine diatom, Skeletonema costatum. Upregulation of ScDSP-1 mRNA was correlated to genomic DNA fragmentation in aging cell populations, which implies its participation in the regulatory mechanisms of S. costatum growth at the molecular level (11). However, it is still unclear what types of environmental stresses cause a response of ScDSP-1. The roles that ScDSP-1 plays in the molecular pathway leading to PCD in diatoms are equally unclear. In this study, we present evidence to demonstrate that blockage of electron flow in photosynthesis, due to either darkness or photosynthesis inhibitors, is the major cause for an increase in the ScDSP-1 transcript level. Furthermore, we conducted experiments to confirm that on the pathway from environmental stress to autolysis, ScDSP-1 expression is associated with the NO production pathway. Possible connections among NO, calcium ions, and ScDSP-1 expression are discussed.

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Culture conditions.
A unialgal culture of Skeletonema costatum strain Kao was grown in sterilized f/2 enriched seawater medium at 20°C under continuous illumination at a light intensity of 145 µmol m^–2 s^–1. To inhibit the growth of bacteria, penicillin and streptomycin (Invitrogen) were added to the medium at final concentrations of 100 U ml^–1 and 100 µg ml^–1, respectively (11, 30). Cell abundance and morphology were monitored by placing 1 ml of algal culture into a Sedgwick-Rafter counting chamber (Hausser Scientific Partnership), and cells were examined with a light microscope (BX60; Olympus) at a magnification of x100 (31). On day 1 or 2 after inoculation, cell populations were growing exponentially, and these cultures were used for all experiments in this study.

Light manipulations.
In the experiment involving treatment with various intensities of irradiance, exponentially growing cultures were divided into four parts, and each was placed under a different light intensity (145, 102, 44, or 15 µmol m^–2 s^–1) created by specific layers of neutral density filters (ND15; Arri GB, London).

Various chemical treatments.
The PS inhibitors 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB) (Sigma-Aldrich), DCMU (Sigma-Aldrich), and paraquat (Riedel-deHaën) were added at final concentrations of 20, 50, and 100 µM, respectively (28, 47, 55). NO provided by the NO donor diethylamine nitric oxide (DEANO) (Sigma-Aldrich) was added at a final concentration of 0.5 mM (62). When needed, either of two NO scavengers, bovine hemoglobin (Sigma-Aldrich) or 2-(4-carboxyphenyl)-4,5-dihydro-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (c-PTIO) (Cayman Chemical), was added to the cultures before treatments with an NO donor or PS inhibitor, at final concentrations of 5 and 100 µM. The pretreatment time for the NO scavengers was 1 h (39, 45, 47). Treatments with PS inhibitors were performed at room temperature under a continuous irradiance of 145 µmol m^–2 s^–1 for 4 h. Treatment with the NO donor, DEANO, was performed under the same incubation conditions. After individual treatments and incubations, cells were harvested for total RNA extraction and in vivo NO detections as described below.

In vivo NO detection.
Exponentially growing S. costatum cells were preincubated in f/2 medium containing a cell-penetrating NO indicator, 4-amino-5-methylamino-2',7'-difluorofluorescein diacetate (DAF-FM diacetate) (Molecular Probes), at a final concentration of 2.5 µM for 30 min (34). Subsequently, cells containing DAF-FM diacetate were washed twice with f/2 medium and then treated with various photosynthesis inhibitors and the NO scavenger, c-PTIO, as described above. Next, cells with a positive signal of green fluorescence were observed with an epifluorescence microscope at excitation wavelengths of 450 to 490 nm and emission wavelengths of 515 nm (Axioplan 2; Zeiss).

Total RNA extraction.
Approximately 10⁷ cells were collected by filtration using a 2-µm-pore polycarbonate membrane (Nuclepore), followed by resuspension of the cell pellet in 0.7 ml guanidine isothiocyanate-containing buffer (RLT buffer; Qiagen) containing 1% β-mercaptoethanol. After disruption of the cells by sonication (sonicator ultrasonic processor XL; Heat System) on ice, total RNA was isolated using the silica membrane spin column included in the RNeasy plant minikit (Qiagen) according to the manufacturer's instructions. The isolated crude RNA was treated with DNase I (RNase free; Roche) at 37°C for 1 h to remove genomic DNA and was subsequently purified by an acidic phenol (pH 4.0)-chloroform-isoamyl alcohol (25:24:1, vol/vol/vol) extraction. The RNA concentration was determined by spectrophotometry (U-2000; Hitachi) at wavelengths of 260 and 280 nm.

Real-time Q-RT-PCR.
DNase I-treated total RNA (1 µg) was reverse transcribed into first-strand cDNA using random hexamers (Promega) and ImpromII reverse transcriptase (Promega) at 25°C for 10 min and 48°C for 1 h. Quantitative PCRs were initiated by adding the cDNA fragments to 1x Sybr green PCR master mix (Applied Biosystems) containing 300 nM of each of the forward and reverse primers. The nucleotide sequences of the primer pair used in the quantitative reverse-transcription PCR (Q-RT-PCR) consisted of ScDSP-SG-F (5'-GAACA AGCAA ACTGC ACTCG TC-3') and ScDSP-SG-R (5'-GTCAA GAATG TTGGT CGTCG CG-3') for ScDSP-1. For the determination of ScDSP-2 mRNA abundance, the primer pair used in the Q-RT-PCR was ScDSP-SG-F and a specific primer, ScDSP-2-SG-R (5'-GTAGG CATCT GCTAT TCTTT CTG-3'). In addition, Ske-18S-F (5'-GAATT CCTAG ATATC GCAGT TCATC-3') and Ske-18S-R (5'-GCTAA TCCAC AATCT CGACT CCTC-3') were used to quantify 18S rRNA. The reactions were then carried out in a GeneAmp 7000 sequence detection system (Applied Biosystems). PCR conditions were set to 95°C for 10 min for 1 cycle followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. The threshold cycle at which the fluorescence intensity exceeded a preset threshold was used to calculate the target gene mRNA and 18S rRNA expression levels. The RNA molar ratio of ScDSP mRNA and 18S rRNA was calculated by the formula described by Chung et al. (11). The specificity of the Q-RT-PCR was confirmed by performing a melting temperature analysis with the GeneAmp 7000 sequence detection system (Applied Biosystems) and was also examined by electrophoresis on a 3% agarose gel containing 0.5x Tris-boric acid-EDTA buffer.

Genomic DNA library construction and screening.
The genomic DNA of S. costatum was isolated by a phenol-chloroform extraction method, and excess polysaccharide was removed with cetyltrimethylammonium bromide (Sigma-Aldrich) (33). After partial digestion with the restriction enzyme Sau3AI, the DNA fragments were fractionated with a 10% to 40% sucrose gradient ultracentrifuge at a speed of 25,000 rpm overnight (SRP28SA; Hitachi). Subsequently, the fraction of DNA fragments ranging from 15 to 20 kb was collected and cloned into the bacteriophage lambda Dash II vector (Stratagene). The S. costatum lambda Dash II genomic DNA library was screened using a 669-bp digoxigenin-labeled ScDSP-1 DNA probe containing the full-length nucleotide sequence of the coding region (11). The washing process was carried out with 2x SSC (1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate)-0.1% sodium dodecyl sulfate and 0.5x SSCi-0.1% sodium dodecyl sulfate at 55°C, and finally CDP-Star (Tropix) was used for chemiluminescent detection. Bacteriophage DNA isolation was conducted as described by Donovan et al. (17).

Identification of DSP cDNA fragments in other marine diatoms.
Thalassiosira pseudonana CCMP 1335 and Phaeodactylum tricornutum CCMP 632 were obtained from the Provasoli-Guillard National Center for Culture of Marine Phytoplankton. Culture conditions, total RNA isolation, and the RT reaction were the same as those described above. The PCR used a reaction mixture containing 3 mM MgCl₂, 200 µM deoxynucleoside triphosphates, 2.5 units of the SuperTaq DNA polymerase (HT Biotechnology), and 500 nM each of the forward and reverse primers. PCR conditions were set to 1 cycle of 95°C for 2 min; 35 cycles of 95°C for 30 s, 62°C for 30 s, and 72°C for 1 min; and then 1 cycle of 72°C for 10 min. The primer pairs used in the PCR were TpDSP1-5N-F (5'-ATGAT TGCTC AA AAG AAAGC CCTC-3') and TpDSP1-3C-R (5'-TTATC TTTAC ACCAA CAATC CCATG-3') for T. pseudonana DSP-1 (TpDSP-1), TpDSP2-5N-F (5'-ATGAT TGCTC CTCAA CGAAA AGCA C-3') and TpDSP2-3C-R (5'-CTACA CCAAC AACCC CAAGT CTC-3') for TpDSP-2; and PtDSP-5N-F (5'-ATGGC CAAGC TTACT TCGAT CGC-3') and PtDSP-3C-R (5'-CTACA CAAGA AAACC CAGGT CAC-3) for Phaeodactylum tricornutum DSP (PtDSP). The PCR product was ligated into the pGEM-T vector (Promega). DNA sequencing and analysis were performed as described below.

DNA and peptide sequence analysis.
The sequence of S. costatum genomic DNA was analyzed using an ABI Prism 377A DNA sequencer with the Prism Ready Reaction Big-Dye termination cycle sequencing kit (Applied Biosystems). The genome information for the other two diatoms, T. pseudonana and P. tricornutum, and the coccolithophorid Emiliania huxleyi were obtained from the DOE Joint Genome Institute (http://www.jgi.doe.gov/). The nucleic acid and deduced amino acid sequences were analyzed using Lasergene software (DNASTAR). Exons and introns in genomic DNA were identified by GENSCAN software (http://genes.mit.edu/GENSCAN.html) and also by comparing genomic DNA sequences with corresponding cDNA sequences. Both BLASTX and BLASTP algorithms from the National Center for Biotechnology Information website (http://www.ncbi.nlm.nih.gov) were also used for sequence analysis.

Nucleotide sequence accession numbers.
The nucleotide sequences of ScDSP-2, PtDSP, TpDSP-1, and TpDSP-2 have been deposited in the GenBank database under accession numbers EF590267, EF590270, EF590268, and EF590269, respectively.

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DSP genes in marine diatoms.
To elucidate the gene structure of ScDSP-1, a Skeletonema costatum genomic DNA library was screened with a probe containing the entire nucleotide sequence of the ScDSP-1 coding region, and accordingly, one positive clone was obtained after screening about 2 x 10⁷ plaques. In this scaffold, the gene structure of ScDSP-1 was revealed to contain a single exon with no introns. Interestingly, another gene, named ScDSP-2, was concatenated at the upstream position 613 bp from the ScDSP-1 start codon, and the putative amino acid sequence of ScDSP-2 shared a high identity of 73.7% with that of ScDSP-1. In addition, the entire genomic sequences of two other marine diatoms, Thalassiosira pseudonana and Phaeodactylum tricornutum, and one coccolithophorid, Emiliania huxleyi, were searched for DSP orthologues (DOE Joint Genome Institute; http://www.jgi.doe.gov/). In T. pseudonana, the concatenation of two DSP-orthologous genes, named TpDSP-1 and TpDSP-2, was found in the position of chr_19a_19:bp 186400 to 188100 (version 3 of T. pseudonana genome annotation). In contrast, in P. tricornutum, one DSP-orthologous gene existed in the position of chr_14:bp 39896 to 399130 (version 2 of P. tricornutum genome annotation). The presence of these DSP orthologues, TpDSP-1, TpDSP-2, and PtDSP, in T. pseudonana and P. tricornutum was further confirmed by RT-PCR. Similar to ScDSP-1 and ScDSP-2, the DSP orthologues in T. pseudonana and P. tricornutum consisted of a single exon. Moreover, a DSP-orthologue gene, E. huxleyi DSP (EhDSP), was found in the position of scaffold 4:bp 1290159 to 1290768 in E. huxleyi (version 1 of E. huxleyi genome annotation). All putative DSP peptides shared a common structure, with a signal peptide near the N terminus and a pair of EF hand motifs for calcium ion binding near the C terminus (40) (Fig. 1).

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FIG. 1. Alignment of five putative DSP peptide sequences from three diatoms, P. tricornutum, S. costatum, and T. pseudonana, and one coccolithophorid, E. huxleyi. The signal peptide and two calcium binding motifs (EF hand) are denoted above the sequences. The conserved amino acid sequences are in black. The names of the putative peptides are given on the left, and the numbers on the right indicate amino acid positions.

Effects of irradiance on ScDSP-1 expression.
Exponentially growing cells under continuous illumination of 145 µmol m^–2 s^–1 were transferred to several cultures exposed to different light intensities (145, 102, 44, and 15 µmol m^–2 s^–1) to test the effect of light on ScDSP-1 expression. Under the irradiances of 145 and 102 µmol m^–2 s^–1, the cell populations grew vigorously from 1.5 x 10⁴ to 1.2 x 10⁵ cells ml^–1 during the 72-h period after initiation of treatment and remained at the highest abundances until day 6. Under the lower irradiance of 44 µmol m^–2 s^–1, cells grew from 1.5 x 10⁴ to 3.2 x 10⁴ cells ml^–1 during the first 24 h after transfer, and then a decrease in population density occurred during the period from day 1 to 2, which was followed by a mild increase to 4.4 x 10⁴ cells ml^–1 on day 6. At the weakest irradiance of 15 µmol m^–2 s^–1, the cell population showed no obvious growth and remained at a low abundance of 1.5 x 10⁴ cells ml^–1 (Fig. 2A). During the experimental period, ScDSP-1 mRNA levels in the two cultures under the higher irradiances of 145 and 102 µmol m^–2 s^–1 obviously increased from 1.1 to 58.2 and 44.6 µmol (mol 18S rRNA)^–1, respectively, as the culture aged (Fig. 2B and C). Under the lower irradiance of 44 µmol m^–2 s^–1, ScDSP-1 mRNA reached a lower level of 8.9 µmol (mol 18S rRNA)^–1 on day 6. However, a peak of ScDSP-1 mRNA occurred between days 1 and 3, with a local maximum of 4.1 µmol (mol 18S rRNA)^–1 (Fig. 2D). In contrast, ScDSP-1 mRNA concentrations in the culture under the weakest irradiance of 15 µmol m^–2 s^–1 remained at a low level of 0.6 to 1.5 µmol (mol 18S rRNA)^–1, but a peak with a value of 17.2 µmol (mol 18S rRNA)^–1 abruptly appeared between days 2 and 3 (Fig. 2E).

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FIG. 2. Effects of irradiance intensities on population growth (A) and ScDSP-1 mRNA abundances in S. costatum (B to E). When cells entered exponential growth, the culture was divided into four parts. One was cultivated under the original illumination of 145 µE m–2 s–1, and the other three were cultivated under illumination levels of 102, 44, and 15 µE m–2 s–1. Error bars indicate standard errors (n = 4). For data points without an error bar, the error bar was smaller than the symbol.

Effects of photosynthesis inhibitors on ScDSP expression.
Because light intensity appeared to be an effective factor controlling the expression of ScDSP-1, PS inhibitors, including the two electron flow blockers DCMU and DBMIB, and a methyl viologen family herbicide, paraquat, were used to explore the physiological functions of ScDSP-1 in S. costatum. With DCMU treatment, ScDSP-1 mRNA levels dramatically increased from 3.5 to 942 µmol (mol 18S rRNA)^–1 within 1 h and further increased to 1,175 µmol (mol 18S rRNA)^–1 by the third hour of treatment (Fig. 3A). With DBMIB treatment, ScDSP-1 mRNA abundance also showed a noticeable increase from 0.5 to 71.5 µmol (mol 18S rRNA)^–1 in 1 h, followed by a mild decrease to 50.7 µmol (mol 18S rRNA)^–1 by the third hour of treatment (Fig. 3B). Nevertheless, treatment with a high concentration of paraquat at 100 µM did not elicit ScDSP-1 expression. mRNA remained at a low level of around 3.5 µmol (mol 18S rRNA)^–1 during the first 3 h of treatment and showed no obvious increase compared to that in the untreated control (Fig. 3C).

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FIG. 3. Effects of the photosynthesis inhibitors DCMU (50 µM) (A), DBMIB (10 µM) (B), and paraquat (100 µM) (C) on ScDSP-1 mRNA expression. Cultures were kept under continuous light at 145 µmol m–2 s–1. Exponentially growing cultures were divided into two parts; one served as the untreated control (solid bars), and the other was treated with various photosynthesis inhibitors (hatched bars) at hour 0. Error bars indicate standard errors (n = 4). For data points without an error bar, the error bar was too small to be clearly drawn.

Next, to determine the ScDSP-2 mRNA levels in S. costatum, a specific reverse primer, ScDSP-2-SG-R, was designed according to the variable region of ScDSP-2 (amino acids 49 to 55) (Fig. 1). This specific reverse primer was used together with the conserved forward primer, ScDSP-SG-F, in the Q-RT-PCR. The melting temperature of the ScDSP-2 amplicon determined with the GeneAmp 7000 sequence detection system was 81.5°C, which significantly differed from the value of 84°C for the ScDSP-1 amplicon. Regardless of whether or not a culture was treated with DCMU, the ScDSP-2 mRNA abundance was only 0.01% of that of ScDSP-1 (Fig. 4). Therefore, the effective quantification of ScDSP-2 mRNA was supported by its characteristic quantity and melting temperature of the PCR product. However, if ScDSP-2 was considered alone, DCMU still induced expression of this gene. The ScDSP-2 mRNA abundance increased from 1.6 x 10^–4 to 3.2 x 10^–2 µmol (mol 18S rRNA)^–1 after 1 h of DCMU treatment, representing a 200-fold increase (Fig. 4).

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FIG. 4. Comparison of ScDSP-1 (solid bars) and ScDSP-2 (hatched bars) mRNA levels with and without the addition of DCMU. Cultures were kept under continuous light at 145 µmol m–2 s–1. Exponentially growing cultures were divided into two parts; one served as the untreated control, and the other was treated with DCMU (50 µM) for 1 h. Error bars indicate standard errors (n = 4). For data points without an error bar, the error bar was too small to clearly be drawn.

Effects of NO on ScDSP-1 expression.
Treatment with the NO donor DEANO dramatically increased ScDSP-1 mRNA abundance from 55.1 µmol (mol 18S rRNA)^–1 to the highest level of 1,218 µmol (mol 18S rRNA)^–1 after 4 h of treatment. Interestingly, this inductive effect of NO was significantly suppressed to give a low level of 297 µmol (mol 18S rRNA)^–1 in the presence of the NO scavenger hemoglobin (Fig. 5).

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FIG. 5. Effect of NO on ScDSP-1 mRNA abundances. Cultures were kept under continuous light at 145 µmol m–2 s–1. Exponentially growing cultures were divided into four parts; one was the untreated control, and the other three parts treated with hemoglobin (Hb) (5 µM), DEANO (0.5 mM), and DEANO plus Hb (D+Hb) for 4 h. One-way analysis of variance and Fisher's least-significant-difference method were conducted among differently treated samples. Different letters indicate a statistically significant difference (P < 0.01). Error bars indicate standard errors (n = 4).

To further confirm that NO is the key messenger that upregulates ScDSP-1 expression when photosynthesis is damaged, another cell-penetrating NO scavenger, c-PTIO, was used in the next assay. At the same time, intracellular NO levels were monitored with an in vivo NO-specific indicator, DAF-FM diacetate. As in the previous experiment, treatment with DCMU or DBMIB elevated ScDSP-1 mRNA levels in S. costatum (Fig. 4 and 6), and the magnitudes of enhancement were in accordance with the increases in NO-producing cells. The enhancement due to the addition of DCMU was effectively alleviated by pretreatment with c-PTIO. ScDSP-1 mRNA levels in c-PTIO-pretreated cultures dropped substantially from 1,670 to 375 µmol (mol 18S rRNA)^–1, and the fraction of NO-producing cells decreased from 62.6% to 29.3% (Fig. 6). In DBMIB-treated cultures, while pretreatment with c-PTIO also resulted in the reduction of ScDSP-1 mRNA abundance, it did not reduce the fraction of NO-producing cells (Fig. 6). With paraquat treatment, there were no differences in either the fraction of NO-producing cells or ScDSP-1 mRNA levels compared to those in the untreated control (Fig. 6).

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FIG. 6. Effects of pretreatment with the cell-penetrating NO scavenger c-PTIO on the percentage of NO-producing cells (A) and ScDSP-1 mRNA levels (B) of S. costatum caused by different PS inhibitors. Cultures were kept under continuous light at 145 µmol m–2 s–1. Exponentially growing cultures were pretreated with c-PTIO for 1 h and then treated with different photosynthesis inhibitors for another hour. (A) Proportion of cells with a positive signal for internal NO as determined by epifluorescence microscopy (with about 500 cells examined per sample). (B) ScDSP-1 mRNA levels. Solid bars, samples treated with individual inhibitors only; hatched bars, samples pretreated with c-PTIO, followed by treatment with individual inhibitors. One-way analysis of variance and Fisher's least-significant-difference method were conducted among different treatments. Different letters indicate a statistically significant difference (P < 0.05). Error bars indicate standard errors (n = 5 for NO-producing cell counts; n = 4 for ScDSP-1 mRNA abundance measurements).

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A survey of decoded genomic databases for several species of phytoplankton was carried out, and orthologous genes encoding DSPs were found exclusively in the genomes of two marine diatoms, Thalassiosira pseudonana and Phaeodactylum tricornutum, and in the coccolithophorid Emiliania huxleyi (Fig. 1). According to the origins of plastids, diatoms and coccolithophorids are descended from the "red alga" branch in the evolution of photosynthetic organisms (22, 41). On the other hand, no DSP-orthologous genes were found in a search of genome databases of phytoplankton belonging to the "green alga" branch, including Chlamydomonas reinhardtii, Ostreococcus lucimarinus, and Ostreococcus tauri. This evolutionary divergence yielded many unique genes in diatoms, which led to the fact that the physiological functions of about half of the genes in the diatom genome cannot be assigned on the basis of similarities to known genes in other organisms (2). Our results based on genome searches suggested that DSPs exist solely in cells belonging to the "red alga" branch.

It has previously been shown that among several stressful conditions tested, including light deprivation, nutrient starvation, and unfavorable temperatures, an increase in ScDSP-1 mRNA occurred only in exponentially growing cells cultivated under a low irradiance of 15 µmol m^–2 s^–1 (10). In the current study, results from both the light manipulation experiment and the photosynthesis inhibitor experiments indicated an intimate relationship between ScDSP expression and photosynthesis (Fig. 2, 3, and 4). In the literature, two positions in the light reaction of photosynthesis have been suggested to be the checkpoints regulating both plastid and nuclear gene expression. One is the plastoquinone in PS-II, and the other is the Q_o site in the cytochrome b₆f complex (4, 18, 43, 44, 51, 60, 61). Since upregulation of ScDSP mRNA occurred in cultures treated with either DCMU or DBMIB but did not occur in cultures treated with paraquat (Fig. 3), the expression of ScDSP was probably controlled at the Q_o site of the cytochrome b₆f complex. This did not exclude the existence of checkpoints, however. In addition, the ScDSP-2 mRNA level was negligibly small compared to that of ScDSP-1 (Fig. 4). The possibility that ScDSP-2 may participate in a different physiological pathway needed to be explored.

If blocking electron flow by DCMU or DBMIB causes a dramatic increase in ScDSP-1 mRNA, then it is quite reasonable to see a similar response in S. costatum cultures transferred from high light to low light in which the electron flow should be greatly reduced (Fig. 2). In the latter case, an important difference is that the upregulation of ScDSP-1 by dim light was transient, suggesting that the expression of ScDSP-1 was no longer required after cells had acclimated to the new light conditions. In unicellular algae, a number of cellular characteristics change in response to variations in environmental light levels. Both the chlorophyll content and the ratio between PS-II and PS-I reaction centers greatly increased in S. costatum cultivated under a low irradiance of around 20 µE m^–2 s^–1 (19, 20). Additionally, increases in the chloroplast volume and thylakoid surface density were observed in another diatom, Cyclotella meneghiniana, grown under low irradiance (46). As a result, the sudden appearance of the transient ScDSP-1 mRNA peak under dim light suggests that ScDSP-1 is an upstream molecule in the light-shade acclimation process in marine diatoms (Fig. 2D and E).

Under high light conditions, ScDSP-1 mRNA did not remain at a low level throughout the entire experimental period. Instead, it rapidly increased with the age of the culture (Fig. 2B and C). In the stationary phase, since the cessation of population growth might have been caused by various combinations of insufficient light, nutrient starvation, and self-generated wastes, it is uncertain if the blockage of electron flow is the only mechanism which can stimulate the expression of ScDSP-1. However, a useful clue is the cooccurrence of PCD and NO in aged S. costatum cultures (11, 64). NO serves as a comprehensive molecule for signal transduction pathways when organisms are experiencing abiotic or biotic stresses (12, 15, 29). In higher plants, NO was identified as being an important messenger in defense responses and growth regulation (14, 32, 59). Using a proteomics approach, a set of photosynthesis-related proteins regulated by NO was identified in the leaves of mung bean (Phaseolus aureus) (37). According to our results, the increase in ScDSP-1 mRNA levels was highly responsive to the generation of NO (Fig. 5). In a previous study, we reported that ScDSP-1 mRNA levels increased with culture age (11). It is also known that NO concentrations in S. costatum culture reach the highest level at the end of exponential growth (64). Together, these lines of evidence strongly suggest that the age-dependent expression of ScDSP-1 in S. costatum is mediated by NO signaling. As for the cause of elevated intracellular NO, studies with several unicellular algae, including C. reinhardtii, Scenedesmus obliquus, and Anabaena doliolum, all indicated that the blocking of PS-II electron transport due to a lack of light or DCMU treatment was responsible (38, 47).

In higher plants and algae, two routes are known for the production of intracellular NO (7, 16). The NO synthase pathway has been well characterized in animals, and several NO synthases were recently identified in Arabidopsis thaliana (32, 59). An analysis of the complete genomes of two diatoms, T. pseudonana and P. tricornutum, revealed that orthologues of NO synthases do exist in diatom cells (1). In addition, it has been reported that NO generated from a calcium-dependent NO synthase is used by the diatom P. tricornutum as a means of chemical defense against copepod grazing (57). Alternatively, intracellular NO can be generated via the nitrate reductase pathway under conditions of light deprivation and malfunction of cellular metabolism (27, 47, 52, 62, 63). Since the addition of the NO synthase inhibitor N-nitro-L-arginine failed to decrease the mRNA level of ScDSP-1 in DCMU-treated S. costatum (C.-C. Chung, unpublished result), we speculated that the source of intracellular NO was unlikely to be the NO synthase pathway when the PS-II electron flow was blocked. Of course, our inference does not exclude the possibility that ScDSP-1 may respond to NO generated via the NO synthase pathway under conditions other than PS-II blockage.

Based on these discussions, the blockage of electron flow between PS-II and cytochrome b₆f was the main cause for the elevated NO production, which subsequently resulted in an increase in ScDSP-1 mRNA. In addition to NO, the generation of ROS is another process considered to be important in terminating algal blooms (35, 56, 58). It is reasonable to speculate that ROS also participate in the regulatory pathway of ScDSP-1. This question is currently under study and will be elucidated in the future.

 
We thank the Core Facility of the Institute of Cellular and Organismic Biology, Academia Sinica, for assistance with DNA sequencing.

This study was supported by grants NSC94-2313-B-019-030 and NSC94-2611-M-019-002 from the National Science Council and by the Center for Marine Bioscience and Biotechnology, National Taiwan Ocean University.

 
* Corresponding author. Mailing address: Institute of Marine Biology, National Taiwan Ocean University, Keelung 20224, Taiwan, Republic of China. Phone: 886-2-2462-2192, ext. 5308. Fax: 886-2-2463-3152. E-mail: jengchang{at}mail.ntou.edu.tw

Published ahead of print on 5 September 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Allen, A. E., A. Vardi, and C. Bowler. 2006. An ecological and evolutionary context for integrated nitrogen metabolism and related signaling pathways in marine diatoms. Curr. Opin. Plant Biol. 9:264-273.[CrossRef][Medline]
2
2. Armbrust, E. V., J. A. Berges, C. Bowler, B. R. Green, D. Martinez, N. H. Putnam, S. Zhou, A. E. Allen, K. E. Apt, M. Bechner, M. A. Brzezinski, B. K. Chaal, A. Chiovitti, A. K. Davis, M. S. Demarest, J. C. Detter, T. Glavina, D. Goodstein, M. Z. Hadi, U. Hellsten, M. Hildebrand, B. D. Jenkins, J. Jurka, V. V. Kapitonov, N. Kroger, W. W. Y. Lau, T. W. Lane, F. W. Larimer, J. C. Lippmeier, S. Lucas, M. Medina, A. Montsant, M. Obornik, M. S. Parker, B. Palenik, G. J. Pazour, P. M. Richardson, T. A. Rynearson, M. A. Saito, D. C. Schwartz, K. Thamatrakoln, K. Valentin, A. Vardi, F. P. Wilkerson, and D. S. Rokhsar. 2004. The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and metabolism. Science 306:79-86.[Abstract/Free Full Text]
3
3. Asada, K. 2006. Production and scavenging of reactive oxygen species in chloroplasts and their functions. Plant Physiol. 141:391-396.[Free Full Text]
4
4. Baier, M., and K.-J. Dietz. 2005. Chloroplasts as source and target of cellular redox regulation: a discussion on chloroplast redox signals in the context of plant physiology. J. Exp. Bot. 56:1449-1462.[Abstract/Free Full Text]
5
5. Berges, J. A., and P. G. Falkowski. 1998. Physiological stress and cell death in marine phytoplankton: induction of proteases in response to nitrogen or light limitation. Limnol. Oceanogr. 43:129-135.
6
6. Berman-Frank, I., K. D. Bidle, L. Haramaty, and P. G. Falkowski. 2004. The demise of the marine cyanobacterium, Trichodesmium spp., via an autocatalyzed cell death pathway. Limnol. Oceanogr. 49:997-1005.
7
7. Besson-Bard, A., A. Pugin, and D. Wendehenne. 2008. New insights into nitric oxide signaling in plants. Annu. Rev. Plant Biol. 59:21-39.[CrossRef][Medline]
8
8. Bidle, K. D., and P. G. Falkowski. 2004. Cell death in planktonic, photosynthetic microorganisms. Nat. Rev. Microbiol. 2:643-655.[CrossRef][Medline]
9
9. Brussaard, C. P. D., R. Riegman, A. A. M. Noordeloos, G. C. Cadée, H. Witte, A. J. Kop, G. Nieuwland, F. C. van Duyl, and R. P. M. Bak. 1995. Effects of grazing, sedimentation and phytoplankton cell lysis on the structure of a coastal pelagic food web. Mar. Ecol. Prog. Ser. 123:259-271.[CrossRef]
10
10. Chung, C.-C. 2004. The identification of growth-dependent genes and their potential application to the study of marine phytoplankton ecology. Ph.D. thesis. National Taiwan Ocean University, Keelung, Taiwan.
11
11. Chung, C.-C., S.-P. L. Hwang, and J. Chang. 2005. Cooccurrence of ScDSP gene expression, cell death, and DNA fragmentation in a marine diatom, Skeletonema costatum. Appl. Environ. Microbiol. 71:8744-8751.[Abstract/Free Full Text]
12
12. Crawford, N. M., and F.-Q. Guo. 2005. New insights into nitric oxide metabolism and regulatory functions. Trends Plant Sci. 10:195-200.[CrossRef][Medline]
13
13. Damste, J. S. S., G. Muyzer, B. Abbas, S. W. Rampen, G. Masse, W. G. Allard, S. T. Belt, J.-M. Robert, S. J. Rowland, J. M. Moldowan, S. M. Barbanti, F. J. Fago, P. Denisevich, J. Dahl, L. A. F. Trindade, and S. Schouten. 2004. The rise of the rhizosolenid diatoms. Science 304:584-587.[Abstract/Free Full Text]
14
14. Delledonne, M., Y. Xia, R. A. Dixon, and C. Lamb. 1998. Nitric oxide functions as a signal in plant disease resistance. Nature 394:585-588.[CrossRef][Medline]
15
15. Delledonne, M. 2005. NO news is good news for plants. Curr. Opin. Plant Biol. 8:390-396.[CrossRef][Medline]
16
16. del Río, L. A., J. F. Corpas, and J. B. Barroso. 2004. Nitric oxide and nitric oxide synthase activity in plants. Phytochemistry 65:783-792.[CrossRef][Medline]
17
17. Donovan, J., X. Lu, and L. Nagarajan. 1993. Rapid purification of bacteriophage DNA. BioTechniques 15:602-603.[Medline]
18
18. Escoubas, J., M. Lomas, J. LaRoche, and P. G. Falkowski. 1995. Light intensity regulation of cab gene transcription is signaled by the redox state of the plastoquinone pool. Proc. Natl. Acad. Sci. USA 92:10237-10241.[Abstract/Free Full Text]
19
19. Falkowski, P. G., and T. G. Owens. 1980. Light-shade adaptation. Plant Physiol. 66:592-595.[Abstract/Free Full Text]
20
20. Falkowski, P. G., T. G. Owens, A. C. Ley, and D. C. Mauzerall. 1981. Effects of growth irradiance levels on the ratio of reaction centers in two species of marine phytoplankton. Plant Physiol. 68:969-973.[Abstract/Free Full Text]
21
21. Falkowski, P. G., R. T. Barber, and V. Smetacek. 1998. Biogeochemical controls and feedbacks on ocean primary production. Science 281:200-206.[Abstract/Free Full Text]
22
22. Falkowski, P. G., M. E. Katz, A. H. Knoll, A. Quigg, J. A. Raven, O. Schofield, and F. J. R. Taylor. 2004. The evolution of modern eukaryotic phytoplankton. Science 305:354-360.[Abstract/Free Full Text]
23
23. Field, C. B., M. J. Behrenfeld, J. T. Randerson, and P. G. Falkowski. 1998. Primary production of the biosphere: integrating terrestrial and oceanic components. Science 281:237-240.[Abstract/Free Full Text]
24
24. Franklin, D. J., C. P. D. Brussaard, and J. A. Berges. 2006. What is the role and nature of programmed cell death in phytoplankton ecology? Eur. J. Phycol. 41:1-14.[CrossRef]
25
25. Fuhrman, J. A. 1999. Marine viruses and their biogeochemical and ecological effects. Nature 399:541-548.[CrossRef][Medline]
26
26. Gechev, T. S., F. van Breusegem, J. M. Stone, I. Denev, and C. Laloi. 2006. Reactive oxygen species as signals that modulate plant stress responses and programmed cell death. BioEssays 28:1091-1101.[CrossRef][Medline]
27
27. Giordano, M., Y.-B. Chen, M. Koblizek, and P. G. Falkowski. 2005. Regulation of nitrate reductase in Chlamydomonas reinhardtii by the redox state of the plastoquinone pool. Eur. J. Phycol. 40:345-352.[CrossRef]
28
28. Goyal, A., and N. E. Tolbert. 1996. Association of glycolate oxidation with photosynthetic electron transport in plant and algal chloroplasts. Proc. Natl. Acad. Sci. USA 93:3319-3324.[Abstract/Free Full Text]
29
29. Grün, S., C. Lindermayr, S. Sell, and J. Durner. 2006. Nitric oxide and gene regulation in plants. J. Exp. Bot. 57:507-516.[Abstract/Free Full Text]
30
30. Guillard, R. R. L., and J. H. Ryther. 1962. Studies on marine planktonic diatom. I. Cyclotella nana Hustedt and Detonula confervacae (Cleve) Gran. Can. J. Microbiol. 8:229-239.[Medline]
31
31. Guillard, R. R. L. 1973. Division rates, p. 289-311. In J. R. Stein (ed.), Handbook of phycological methods: culture methods and growth measurements. Cambridge University Press, Cambridge, United Kingdom.
32
32. Guo, F.-Q., M. Okamoto, and N. M. Crawford. 2003. Identification of a plant nitric oxide synthase gene involved in hormonal signaling. Science 302:100-103.[Abstract/Free Full Text]
33
33. Hwang, S.-P. L., S. K. Wang, S. F. Wei, L.-C. Cheng, and J. Chang. 1999. Identification and expression pattern of DNA polymerase gene in a marine diatom, Skeletonema costatum. Mar. Biotechnol. 1:200-206.[CrossRef][Medline]
34
34. Itoh, Y., F. H. Ma, H. Hoshi, M. Oka, K. Noda, Y. Ukai, H. Kojima, T. Nagano, and N. Toda. 2000. Determination and bioimaging method for nitric oxide in biological specimens by diaminofluorescein fluorometry. Anal. Biochem. 287:203-209.[CrossRef][Medline]
35
35. Kahl, L. A., A. Vardi, and O. Schofield. 2008. Effects of phytoplankton physiology on export flux. Mar. Ecol. Prog. Ser. 354:3-19.[CrossRef]
36
36. Koonin, E. V., and L. Aravind. 2002. Origin and evolution of eukaryotic apoptosis: the bacterial connection. Cell Death Differ. 9:394-404.[CrossRef][Medline]
37
37. Lum, H.-K., C.-H. Lee, Y. K.-C. Butt, and S. C.-L. Lo. 2005. Sodium nitroprusside affects the level of photosynthetic enzymes and glucose metabolism in Phaseolus aureus (mung bean). Nitric Oxide 12:220-230.[CrossRef][Medline]
38
38. Mallick, N., L. C. Rai, F. H. Mohn, and C. J. Soeder. 1999. Studies on nitric oxide (NO) formation by the green alga Scenedesmus obliquus and the diazotrophic cyanobacterium Anabaena doliolum. Chemosphere 39:1601-1610.[Medline]
39
39. Mallick, N., F. H. Mohn, C. J. Soeder, and J. U. Grobbelaar. 2002. Ameliorative role of nitric oxide on H₂O₂ toxicity to a chlorophycean alga Scenedesmus obliquus. J. Gen. Appl. Microbiol. 48:1-7.[CrossRef][Medline]
40
40. Nakayama, S., and R. H. Kretsinger. 1994. Evolution of the EF-hand family of proteins. Annu. Rev. Biophys. Biomol. Struct. 23:473-507.[Medline]
41
41. Nisbet, R. E. R., O. Kilian, and G. I. McFadden. 2004. Diatom genomics: genetic acquisitions and mergers. Curr. Biol. 14:R1048-R1050.[CrossRef][Medline]
42
42. Nishiyama, Y., S. I. Allakhverdiev, and N. Murata. 2006. A new paradigm for the action of reactive oxygen species in the photoinhibition of photosystem II. Biochim. Biophys. Acta 1757:742-749.[Medline]
43
43. Pfannschmidt, T., K. Schütze, M. Brost, and R. Oelmüller. 2001. A novel mechanism of nuclear photosynthesis gene regulation by redox signals from the chloroplast during photosystem stoichiometry adjustment. J. Biol. Chem. 276:36125-36130.[Abstract/Free Full Text]
44
44. Pfannschmidt, T. 2003. Chloroplast redox signals: how photosynthesis controls its own genes. Trends Plant Sci. 8:33-41.[CrossRef][Medline]
45
45. Pfeiffer, S., E. Leopold, B. Hemmens, K. Schmidt, E. R. Werner, and B. Mayer. 1997. Interference of carboxy-PTIO with nitric oxide- and peroxynitrite-mediated reactions. Free Radic. Biol. Med. 22:787-794.[CrossRef][Medline]
46
46. Rosen, B. H., and R. L. Lowe. 1984. Physiological and ultrastructural responses of Cyclotella meneghiniana (Bacillariophyta) to light intensity and nutrient limitation. J. Phycol. 20:173-183.[CrossRef]
47
47. Sakihama, Y., S. Nakamura, and H. Yamasaki. 2002. Nitric oxide production mediated by nitrate reductase in the green alga Chlamydomonas reinhardtii: an alternative NO production pathway in photosynthetic organisms. Plant Cell Physiol. 43:290-297.[Abstract/Free Full Text]
48
48. Samuilov, V. D., E. M. Lagunova, D. B. Kiselevsky, E. V. Dzyubinskaya, Y. V. Makarova, and M. V. Gusev. 2003. Participation of chloroplasts in plant apoptosis. Biosci. Rep. 23:103-117.[CrossRef][Medline]
49
49. Saviani, E. E., C. H. Orsi, J. F. P. Oliveira, C. A. F. Pinto-Maglio, and I. Salgado. 2002. Participation of the mitochondrial permeability transition pore in nitric oxide-induced plant cell death. FEBS Lett. 510:136-140.[CrossRef][Medline]
50
50. Segovia, M., L. Haramaty, J. A. Berges, and P. G. Falkowski. 2003. Cell death in the unicellular chlorophyte Dunaliella tertiolecta. A hypothesis on the evolution of apoptosis in higher plants and metazoans. Plant Physiol. 132:99-105.[Abstract/Free Full Text]
51
51. Shao, N., O. Vallon, R. Dent, K. K. Niyogi, and C. F. Beck. 2006. Defects in the cytochrome b₆/f complex prevent light-induced expression of nuclear genes involved in chlorophyll biosynthesis. Plant Physiol. 141:1128-1137.[Abstract/Free Full Text]
52
52. Takahashi, S., and H. Yamasaki. 2002. Reversible inhibition of photophosphorylation in chloroplasts by nitric oxide. FEBS Lett. 512:145-148.[CrossRef][Medline]
53
53. Thornberry, N. A., and Y. Lazebnik. 1998. Caspases: enemies within. Science 281:1312-1316.[Abstract/Free Full Text]
54
54. Tréguer, P., D. M. Nelson, A. J. van Bennekom, D. J. deMaster, A. Leynaert, and B. Queguiner. 1995. The silica balance in the world ocean: a reestimate. Science 268:375-379.[Abstract/Free Full Text]
55
55. Tsuji, N., N. Hirayanagi, M. Okada, H. Miyasaka, K. Hirata, M. H. Zenk, and K. Miyamotoa. 2002. Enhancement of tolerance to heavy metals and oxidative stress in Dunaliella tertiolecta by Zn-induced phytochelatin synthesis. Biochem. Biophys. Res. Commun. 293:653-659.[CrossRef][Medline]
56
56. Vardi, A., I. Berman-Frank, T. Rozenberg, O. Hadas, A. Kaplan, and A. Levine. 1999. Programmed cell death of the dinoflagellate Peridinium gatunense is mediated by CO₂ limitation and oxidative stress. Curr. Biol. 9:1061-1064.[CrossRef][Medline]
57
57. Vardi, A., F. Formiggini, R. Casotti, A. D. Martino, F. Ribalet, A. Miralto, and C. Bowler. 2006. A stress surveillance system based on calcium and nitric oxide in marine diatoms. PLoS Biol. 4:411-419.[CrossRef]
58
58. Vardi, A., D. Eisenstadt, O. Murik, I. Berman-Frank, T. Zohary, A. Levine, and A. Kaplan. 2007. Synchronization of cell death in a dinoflagellate population is mediated by an excreted thiol protease. Environ. Microbiol. 9:360-369.[CrossRef][Medline]
59
59. Wendehenne, D., J. Durner, and D. F. Klessig. 2004. Nitric oxide: a new player in plant signaling and defense responses. Curr. Opin. Plant Biol. 7:449-455.[CrossRef][Medline]
60
60. Wilson, K. E., A. G. Ivanov, G. Öquist, B. Grodzinski, F. Sarhan, and N. P. A. Huner. 2006. Energy balance, organellar redox status, and acclimation to environmental stress. Can. J. Bot. 84:1355-1370.[CrossRef]
61
61. Wollman, F.-A. 2001. State transitions reveal the dynamics and flexibility of the photosynthetic apparatus. EMBO J. 20:3623-3630.[CrossRef][Medline]
62
62. Yamasaki, H., Y. Sakihama, and S. Takahashi. 1999. An alternative pathway for nitric oxide production in plants: new features of an old enzyme. Trends Plant Sci. 4:128-129.[CrossRef][Medline]
63
63. Yamasaki, H., and Y. Sakihama. 2000. Simultaneous production of nitric oxide and peroxynitrite by plant nitrate reductase: in vitro evidence for the NR-dependent formation of active nitrogen species. FEBS Lett. 468:89-92.[CrossRef][Medline]
64
64. Zhang, Z.-B., C.-Y. Liu, Z.-Z. Wu, L. Xing, and P.-F. Li. 2006. Detection of nitric oxide in culture media and studies on nitric oxide formation by marine microalgae. Med. Sci. Monit. 12:BR75-BR85.[Medline]

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Applied and Environmental Microbiology, November 2008, p. 6521-6527, Vol. 74, No. 21
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