Accepted Manuscript Title: Nuclear and Cell Morphological Changes during the Cell Cycle and Growth of the Toxic Dinoflagellate Alexandrium minutumCell Cycle Analysis of Axeandrium minutum–> Author: Carlos Dapena Isabel Bravo Angeles Cuadrado Rosa Isabel Figueroa PII: DOI: Reference:
S1434-4610(15)00002-4 http://dx.doi.org/doi:10.1016/j.protis.2015.01.001 PROTIS 25472
To appear in: Received date: Revised date: Accepted date:
7-7-2014 2-12-2014 8-1-2015
Please cite this article as: Dapena, C., Bravo, I., Cuadrado, A., Figueroa, R.I.,Nuclear and Cell Morphological Changes during the Cell Cycle and Growth of the Toxic Dinoflagellate Alexandrium minutumCell Cycle Analysis of Axeandrium minutum–>, Protist (2015), http://dx.doi.org/10.1016/j.protis.2015.01.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
ORIGINAL PAPER
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Nuclear and Cell Morphological Changes during the Cell Cycle and Growth of the Toxic Dinoflagellate Alexandrium minutum
Instituto Español de Oceanografía (IEO), Subida a Radio Faro 50, 36390 Vigo, Spain
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Carlos Dapenaa, Isabel Bravoa, Angeles Cuadradob; and Rosa Isabel Figueroaa,c,1
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Aquatic Ecology, Biology Building, Lund University, 22362 Lund, Sweden
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Universidad de Alcalá (UAH), Dpto de Biomedicina y Biotecnologia, 28801 Alcalá de Henares, Spain
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Submitted July 7, 2014; Accepted January 8, 2015
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Monitoring Editor: Marina Montresor
Running title: Cell Cycle Analysis of Axeandrium minutum
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Corresponding author; e-mail [email protected]/ [email protected]. (R. I. Figueroa).
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Elucidation of the cell cycle of dinoflagellates is essential to understand the processes leading to their massive proliferations, known as harmful algal blooms. In this study, we used imaging flow cytometry (IFC) to monitor the changes in DNA content and nuclear and cell morphology that occur during clonal growth of the toxic species Alexandrium minutum Halim. Our results indicate that the population was in S phase (C→2C DNA content) during
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the light period, whereas haploid cells with a C DNA content peaked only during a short interval of the dark period. The timing of the phases, identified based on the nuclear
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morphology and cytoplasmic-to-nuclear (CNR) ratio of the cells, suggests that the length of the G2/M phase is regulated by nutrient levels whereas the beginning of S phase is clock
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controlled. In addition we found that up to 7% of individual cells achieved a DNA content higher than 2C, indicative of either zygote formation and replication (homothallism), or of double-haploid cells able to divide (polyploid forms). Cells belonging to different cell cycle
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phases (G1-S-G2/M) could be readily discriminated based on nuclear size. Our study provides evidence of cell-cycle plasticity during clonal growth and unambiguously characterizes the
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cell-cycle phases of this dinoflagellate species.
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Introduction
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Key words: Cell-cycle; cytoplasmic-to-nuclear ratio (CNR); dinoflagellates; growth; nuclear size.
During the cell cycle, eukaryotic cells undergo asexual division to yield two genetically identical daughter cells. The cell cycle proceeds in three stages: i) interphase, marked by three distinct phases: G1, in which growing cells exhibit high metabolic activity; S, during which cellular DNA is replicated; and G2, a temporal gap between S phase and the following division in which cells accumulate the nutrients needed for the division process; ii) mitosis (M phase), during which there is nuclear division, or karyokinesis; and iii) cytokinesis, when cytoplasmic division marks the completion of cell division and thus of the cell cycle. The molecular events that determine a cell's progress through the cell cycle are ordered and directional, such that the cycle is irreversible. There is also an additional stage, called G0, composed of cells that have exited the cycle and stopped dividing. These quiescent cells in Go cannot be differentiated from those in G1, either morphologically or based on their DNA content. However, G0 cells can be activated to re-enter the cell cycle at G1 (see, for example, the review by Pardee 1989). The transition 2
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between different phases of the cell cycle is controlled by cyclins and cyclin-dependent kinases (CDKs), which are highly conserved among eukaryotes. In concert with other species-specific components, cyclins and CDKs regulate the cell cycle itself. Aside from the recognition of the various phases of the cell cycle, eukaryotic diversity in cell-cycle control remains largely unexplored, in part because most of the model species have been unikonts (Harashima et al.
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2013).
The cell cycle is composed of two cycles: the DNA division cycle and the growth cycle,
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which includes the synthesis of all other macromolecules (Vaulot et al. 1986). The DNA
synthesis pattern of many phytoplankton species suggests that a circadian clock controls the
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timing of cell division, with light- and dark-sensitive periods (Chisholm 1981, Chisholm and Brand 1981). Dinoflagellates are unicellular protists, with many species forming harmful algal
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blooms (e.g., Hallegraeff 1993; Smayda 1997). Discrimination between dinoflagellate life- and cell-cycle stages is essential to understand the development of dinoflagellate blooms. During dinoflagellate division, cytokinesis occurs during the day, and karyokinesis/mitosis generally at
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night (Hastings and Sweeney 1964). However, Karentz (1983) observations suggest that there is a huge diversity in the group, different dinoflagellate species differing in their division patterns. For example, DNA synthesis and cell division may be in phase or not, being
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hypothesized that they are in phase in populations with division rates of less than one per day,
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whereas in the others (e.g. Gymnodinium nelsoni or Pyrocystis fusiformis), there is segregation of DNA replication and cell division between different L:D cycles. The same study points to that
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G1, G2 and S length could be specifically regulated among species. Supporting this diversity, DNA synthesis and cell division have been described in dinoflagellates either as a continuous process, similar to that in prokaryotes, or as following a typical eukaryotic G1-S-G2-M phasetransition model (Allen et al. 1975; Polikarpov and Tokareva 1970; Spector et al. 1981), albeit with several singular features (Bhaud et al. 2000). The dinoflagellate nucleus, referred to as the dinokaryon (Rizzo 1991), has many unusual characteristics, most notably its very large genome size. Indeed, the dinoflagellate genome is among the largest of all eukaryotic genomes (Hou and Lin 2009). Dinoflagellate chromosomes, organized by means of a cholesteric liquid crystal structure, are visible throughout the cell cycle (Chow et al. 2010, Rill et al. 1989), but in many respects their morphology resembles that seen in other eukaryotes (Figueroa et al. 2014). The chromosomes divide via closed mitosis, with a permanent nuclear envelope and through cytoplasmic channels (Soyer-Gobillard et al. 1999), but they lack detectable histones, despite the fact that recent studies demonstrated the existence of all core histone mRNA sequences (Lin et al. 2010, Roy and Morse 2012). The characteristics of the dinoflagellate nucleus have
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been attributed to the acquisition of prokaryotic genes during evolution, which may also explain the diversity and relatively slow growth rates of dinoflagellates. The latter has been attributed to their having a Rubisco enzyme, which inefficiently distinguishes CO2 from O2 and probably originated from anaerobic protobacteria (Wong and Kwok 2005).
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In the dinoflagellate Lingulodinium polyedrum, S phase, and not mitosis, was gated by the circadian clock (Homma and Hastings 1989; Lidie et al. 2005). Nonetheless, the results of Dagenais-Bellefeuille et al. (2008) in this species are consistent with a circadian clock
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controlling both the G2/M and S phases. Nutrients have been considered as potential modulators of the cell cycle phases, but the length of the G2 phase in phytoplankton growing
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under different nutritional levels remained constant in the study by Olson and Chisholm (1986).
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Blooms of the cosmopolitan and toxin-producing dinoflagellate Alexandrium minutum Halim occur worldwide in shallow systems such as estuaries, bays, lagoons, and harbors (Bravo
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et al. 2010; Garcés et al. 2004; Giacobbe et al. 1996; Touzet et al. 2010). Their negative effects, including production of the toxin that causes paralytic shellfish poisoning, have been welldocumented in humans and in other species. Growth in Alexandrium, including A. minutum, is
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mainly achieved through asexual division (mitosis). Sexual reproduction can be induced by,
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among other factors, phosphate deficiency and apparently involves more than two compatible mating types (complex heterothallism), given that self-fertilization in clonal strains has yet to
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be observed (Figueroa et al. 2007). In dinoflagellate ecology, sexual reproduction has several benefits in addition to the potential for genetic recombination; mainly in those species in which leads to resting cyst formation (see Bravo and Figueroa 2014 and references therein). Resting cysts are benthic forms that are highly resistant to adverse conditions, and they can remain viable in the sediments for up to 100 years (Miyazono et al. 2012). Germination and subsequent divisions (meiosis) of the resting cyst restore its haploid life-stage. Apart from the cyst stage, which in most dinoflagellate species is morphologically very different from planktonic forms, the life- and cell-cycle stages of Alexandrium are in general very similar. Resting cysts are well characterized by the absence of flagella, a double cell wall, and the presence of brownish pigment deposits. However, most characteristics considered useful in distinguishing flagellated stages (vegetative stages, gametes, and mobile zygotes) are in fact misleading. For example, regarding size, gametes are generally considered to be smaller, and zygotes larger than vegetative cells (see the revision by Bravo and Figueroa 2014). However, for A. minutum, gametes with a size similar to that of the population average and smaller than zygotes have been observed (Figueroa et al. 2007). Immediately after their formation, mobile 4
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zygotes possess two trailing flagella, unlike the uniflagellate cells of vegetative stages. However, this characteristic is easily lost by cells under stress or after cell fixation. Therefore, the identification of Alexandrium life and cell cycle stages is often very challenging but essential to understand growth and survival strategies of these organisms in their natural
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habitats. In their study of toxin production during the cell cycle in Alexandrium fundyense, Taroncher-Oldenburg et al. (1997) showed that prolonged darkness arrested the cells in a
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light-dependent part of the cell cycle (G0), which prevented both progression through G1 and division. DNA synthesis (S phase) started just before the beginning of the dark period, reaching
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a maximum that coincided with the middle of G2/M phase. Earlier studies showed that at the G1/S boundary, eukaryotic cells must pass a size threshold, whereas at the G2/M boundary a
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prerequisite for further cell-cycle progression is the completion of DNA synthesis (Elledge 1996). In this study, we asked whether this pattern was also relevant for A. minutum and if cells of this species could be unambiguously characterized according to their cell-cycle-specific
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morphology and their nuclear and cellular sizes. To answer this question and to determine the timing of each cell-cycle phase, we used imaging flow cytometry (IFC) and followed nuclear and cellular morphologies to study a darkness-synchronized clone of the dinoflagellate
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Alexandrium minutum over a period of 53 h.
Results
Growth
During the first light:dark period (28–48 TPS or h post-synchronization) the number of cells doubled, from 3290 ± 42.4 cells·mL-1 to 6313 ± 245.4 cells·mL-1 (0.94 div·day-1). During the second light:dark period, growth was slower, with the population increasing from 6313 ± 245.4 cells·mL-1 to 8735 ± 49.5 cells·mL-1 (0.47 div·day-1). Although the cell cycle was evaluated by flow cytometry for only the first 53 h, growth of the clonal culture was followed until day 8 post-synchronization, reaching over 30,000 cells·mL-1. DNA Content The cells were classified according to their DNA content, as determined by the peaks in the nuclear fluorescence histogram of Figure 1, as C, C→2C, and 2C. At the beginning of the study 5
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(28 h TPS) (Fig. 1A) and under light conditions (08:00–20:00), these histograms showed the presence of one main peak, with a relatively high coefficient of variation (CV), which widened over time. However, as the cells progressed through the dark period (20:00), two additional peaks clearly appeared. The one to the left of the first peak corresponded to G1 (1C DNA content) and the other, at the far right, to 2C (2C/C=1.95). Consequently, the main peak
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observed under light conditions was that of S phase (C→2C). S ll during the dark period, at
04:00, or 44 h TPS in Fig. 1, the G1 peak shifted slightly, towards the position of the S-phase
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peak. The latter was maintained in the second light period, starting at 48 h TPS (Fig. 1), until the dark period (20:00 or 60h TPS), during which time the size of the 2C peak started to
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increase. Note that the main position of the cell population during the second light period (C→ 2C peak) was closer to the 1C position than during the first light period. Additionally, peaks corresponding to 1C and 2C occurred later, 2 h after the onset of the dark period (22:00),
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whereas the decrease in the 2C peak stopped 2 h after the beginning of the third light period, rather than during the dark period, as observed before. The size of the 2C peak was
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significantly smaller in the second than in the first dark period. At the beginning of the third light period, the main peak shifted slightly towards the C→2C peak, as observed before; however the displacement was again less than during the first light period.
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Consistent with the different peak sizes shown in Figure 1, the maximum percentage of
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cells in G1 or S phase or undergoing division differed substantially between the light:dark periods studied (Fig. 2A, B), although the overall tendencies were similar (Fig. 2C): Maximums
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of 2C cells and cells with a >2C DNA content and minimums of C→2C cells were detected during darkness, whereas C→2C cells reached their maximum numbers during the light period. In the first light:dark period, the percentages of cells in S and G2/M were markedly higher than during the following period. This finding was supported by the fact that the best curve fit was obtained with separate plots, i.e., 27–54 h TPS (Fig. 2A) and 55–79 h TPS (Fig. 2B). From the graphs in Figure 2, we estimated that the beginning of S phase was at 07:00 in both cycles. G2/M phase started at 18:00 and lasted until 06:00 in the first cycle but until 09:00 in the second. S phase was also longer (~2 h) in the second cycle. Individual cells with a DNA content >2C were detected in both cycles and their pattern of appearance in the dark period was similar to that of 2C cells, but with a much smaller percentage of cells: maximally 7.7% in the first cycle and 7.5% in the second cycle. Morphology
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Cell and nuclear area. Figure 3 shows the average (N>10,000) cellular (Fig. 3A) and nuclear (Fig. 3B) areas together with the cell cycle data for individual cells. The values corresponding to the different phases did not overlap, with the largest differences between phases recorded at the nuclear level. Cell and nuclear areas were strongly correlated, following a linear relationship in all cell-cycle phases and when data from these phases were pooled (Fig.
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3C). After the peak in 2C cells, the average cell size after division (C) decreased until around
400 μm2 in the first light:dark period but reached close to 300 μm2 in the second (Fig. 3A), in
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which fewer cells underwent division (Fig. 2A-B). This fact is shown by a negative correlation in the size of cells over time, observed for all DNA contents unless in cells>2C (Supplementary
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Material Fig. S2). The cytoplasmic-to-nuclear ratio (CNR) changed over time and according to DNA content (Fig. 4A), trending to a linear correlation (R2=0.514). The CNRs of cells with C DNA content overlapped slightly with those of cells with C→2C, even if the median values differed
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(Fig. 4A). Pairwise comparisons using t tests showed highly significant differences in CNR values between groups with different DNA content (Supplementary Material Table S1)
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Nuclear morphology during the different cell-cycle stages. The nuclei of cells analyzed by IFC showed either a roundish or a very weakly lobulated morphology for those with a C DNA content, but a more typical U-shaped morphology characteristic of the Alexandrium genus for
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those with a C→2C DNA content, indicative of S phase (Fig. 5). Three groups were defined in a
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specific study of 2C cells: “individual,” “in early cytokinesis,” and “in two-cell chains” (Fig. 6A). The nuclear morphologies were consistent with the cellular classification (Fig. 6B), as individual
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cells had 2C either U-shaped or roundish nuclei; cells in early cytokinesis had either two C nuclei or one elongated 2C nucleus; and two-cell chains consisted of two differentiated cells with two physically separated C nuclei. The percentage of individual cells in division was seen as a distinct peak that corresponded to the maximum of 2C cells per cycle (Fig. 7). The IFCbased determination of cells in early cytokinesis indicated the onset of mitosis at a similar time than the maximum value of CNR of 2C cells, which peaked at cell division (Fig. 4B). According to the CNRs and the curve followed by cells in early cytokinesis, mitosis occurred at 21:00– 23:00 (37-39 h post-synchronization in the first cycle and at 22:00–01:00 (62–65 h postsynchronization in the second cycle (Figs 4B and 7). Using more detailed images, taken at higher magnification, together with those obtained by IFC, we reconstructed the cell cycle process. Figure 8A, B shows cells smaller than the minimum size needed to enter S phase. The nuclei of these cells are small and U-shaped, with very shallow concavities (Fig. 8A), and are roundish in appearance depending on the angle of view (Fig. 8B). A 2C stage, with a well-developed U-shaped nucleus, is shown in Figure 8C; a 7
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putative metaphase cell, with well-separated chromosomes, is shown in Figure 8D, and a late metaphase cell, with double-chromatid chromosomes (arrows), in Figure 8E. Our reconstruction of anaphase and telophase is provided in Figure 8F–H, which shows these two processes until the formation of a two-cell chain, a mitotic product that is subsequently broken to restore the individuality of the cells. The cell depicted in Figure 8J has a round nucleus and
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decondensed chromosomes, similar to many of the 2C individual cells seen in Figure 6B. Lastly, based on its size and that of its nucleus, the cell in Figure 8K is probably one with a DNA
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Discussion
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The study of dinoflagellate proliferation has considerable interest because it can lead to the formation of Harmful Algal Blooms (HABs), with adverse effects on public health, seafood safety and aquaculture (Smayda 2002). The rate of division of a given population of cells is
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under tight control of cell cycle, a key element of this study. The present work focuses in the
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cell cycle characterization of the cosmopolitan PST-producing species Alexandrium mintum, in order to determine both the timing of the DNA content changes and how cell growth,
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estimated by cell and nuclear areas, relate to the progression and stages of the cell cycle. DNA Content Changes
Many phytoplankton species show circadian regulation of cell cycle (Chisholm and Brand 1981; Homma and Hastings 1989; Sweeney and Hastings 1958), which exhibit three main characteristics: they are endogenously generated, self-sustaining, and persist under constant conditions, with light vs. dark conditions and temperature among the most well-studied (see the review of McClung 2006 and references therein). The “escape from light hypothesis” (Pittendrigh 1993) posits that organisms gain an advantage by carrying out light-sensitive processes, such as DNA replication, during the dark portion of the diel cycle. However, in some dinoflagellates, such as Lingulodinium polyedrum (Dagenais-Bellefeuille et al. 2008; Homma and Hastings 1989), S phase occurs during the dark period, whereas in other species, such as Amphidinium operculatum (Leighfield and Van Dolah 2001; Van Dolah and Leighfield 1999), Karenia brevis (Van Dolah et al. 2007), and Alexandrium fundyense (Taroncher-Oldenburg et al. 1997) it takes place, at least in part, during the day. The last authors reported that immediately
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after the cells are released from prolonged darkness, they enter a nonproliferation period equivalent to one generation time, as result of arrest in G0. These cells are too small to enter and progress through G1. For that reason, our first sampling was 27 h after darkness release and it showed that in A. minutum S-phase occurs entirely during the light period. Moreover, in the second light:dark period of our study, when only a fraction of the cells in the population
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divided, the average DNA content of the population still occurred within the S phase (C→2C),
during the light period. A peak corresponding to cells with a C DNA content was observed only
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after G2/M, under conditions of darkness. Interference due to RNA content can be ruled out in these observations given the high concentrations of RNAase used in the study to specifically
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avoid this problem. Interestingly, and in contrast to L. polyedrum (Dagenais-Bellefeuille et al. 2008), the timing and length of S and G2 in A. minutum varied with growth, as both phases were shorter in the first cycle, when the growth rate was higher, and longer in the second,
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when there was less growth of the population. DNA replication and mitosis are controlled by specific proteins whose rate of synthesis changes during the cell cycle (Evans et al. 1983; Giard
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et al. 1973), and growth-dependent variations in the durations of S and G2 have been demonstrated in different cell lines, e.g., in yeasts. In that study, Slater el al. (1977) examined the relation between the lengths of the cell-cycle phases and the growth rate of the culture.
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The largest variability was in the length of G1 (10-fold vs. nearly 2-fold in both S and G2). In
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dinoflagellate species with division rates of less than one per day , the cell cycle is said to be ruled by the circadian clock, meaning that there is a temporal window through which cells that
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have reached a minimum size threshold can proceed through the cycle (Elledge 1996). This implies cell-cycle control at the S and G2 phases. Indeed, in our experiment these two phases similarly varied between light:dark periods, as the difference in the duration of either one was approximately 2 h. Cell growth has to be coordinated with the cell division cycle to allow an appropriate response to nutrient availability (Wong and Kwok et al. 2005). Our results show that all cells duplicated during the first light:dark period but the population decreased their growth rate during the following one, indicating a progressive loss of cell cycle synchrony. The maximum division rate achieved in the first light:dark period (A. minutum usually grows at lower rates) is probably not possible to maintain over long periods of time, because metabolic needs and nutrient uptake become imbalanced when synchronization is lost (e.g., TaroncherOldenburg et al. 1997, Vaulot et al. 1996). Supporting this theory, during the 53 h of our study the trend was a decrease in the average area of C cells, which suggests nutritional depletion. Thus, dinoflagellates are apparently able to rapidly adapt to the available nutritional resources, modulating their cell-cycle progression by increasing the length of both G2 and S in response
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to the number of cells in G1 that are able to reach the minimum threshold size needed for entry into S phase. Another implication of our results is related to the validity of the estimations of cellular DNA content in some dinoflagellate species. Those estimates were mainly based on
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unsynchronized cultures sampled during light conditions (e.g., LaJeunesse et al.2005). However, according to the results of our study, the DNA content of some dinoflagellate species may be overestimated by this method when S phase starts immediately before the beginning
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of the light period.
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Additionally, another novel result in the present work was the detection of individual cells with a >2C DNA content. One explanation could be related to the sexual cycle of this species. A. minutum has been described as a heterothallic species (Figueroa et al. 2007), as
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two different clones were shown to be necessary to produce sexual resting cysts. However, that study was unable to determine why some zygotes encyst while others divide. We think
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that the existence of cells with a >2C DNA content in the clonal culture (in which no resting cysts were ever observed) is best explained by low levels of homothallism and zygote division (by mitosis or meiosis), as suggested by the fact that cells with a DNA content >2C show a
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similar trend of 2C cells, with peaks during the dark period. Thus, our study is the first to show
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a circadian rhythm for the division of dinoflagellate cells with a ploidy level different than that of haploid cells. While an alternative explanation is the existence of double haploid cells able
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to divide, as a consequence of a possible polyploidization event, we think that the first hypothesis is more plausible because the percentage of these cells increases dramatically following a sexual cross, suggesting that sexuality and zygote formation underlie the origin of cells forming this population (Figueroa et al., unpubl. observ.). Cell and Nuclear Size during the Cell Cycle Nuclear size and shape are carefully controlled in eukaryotic cells, with aberrations in either leading to altered cellular states such as cancer (see the review by Huber and Gerace 2007 and references therein). Cell division requires an increase in the cell surface area:volume ratio (Oney et al. 2005); indeed, there is a well-established correlation between cell and nuclear size, expressed as the CNR, with changes in DNA content altering cell volume (see, for instance, the work in yeasts by Jorgensen et al. 2007). Early observations formed the basis of the “karyoplasmic” ratio theory, according to which eukaryotic cells actively maintain a fixed ratio of nuclear to cell volume (Cavalier-Smith 2005; Wilson 1925). The validity of this hypothesis has been confirmed in cells with a broad range of genetic backgrounds and placed in a wide
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variety of growth conditions, including those that alter cell volume and DNA content. But to our knowledge ours is the first study to confirm this relationship in dinoflagellates, i.e., the eukaryotes with the highest DNA content (Lin 2011). The CNR is an important indicator of cell stability. For example, for the multifunctional
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protein survivin, an inhibitor of apoptosis and a breast cancer biomarker, the CNR was shown to be an important predictor of outcome in tamoxifen-treated patients (Rexhepaj et al. 2010). The variations in the CNRs in our study were mainly but, as evidenced by the data dispersion,
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not exclusively limited to DNA content, consistent with reports of factors besides the amount of DNA in the nucleus affecting the CNR. For example, nuclear size might also be subject to
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regulation depending on cell-type-specific chromatin condensation (Swanson et al. 1991). The CNRs of A. minutum cells in G1 and S overlapped but increased significantly in cells with a DNA
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content ≥2C.
The morphological discrimination between dinoflagellate life- and cell-cycle stages has
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been difficult if not impossible but it is essential if we are to understand dinoflagellate blooming patterns in the field. Our results show that cell and nuclear area, as well as the CNR, can be used to efficiently distinguish between the cell-cycle stages of the dinoflagellate A.
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minutum. Further studies are necessary to determine whether these relationships are
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maintained between genotypes, along the growth curve, and under different environmental conditions, and to design parameters able to accurately evaluate the growth of a bloom and
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forecast its development potential.
Methods
Cell proliferation conditions: The dinoflagellate strain used in this study was A. minutum strain VGO577, regularly maintained at the Centro Oceanográfico de Vigo (CCVIEO; Culture Collection of Harmful Microalgae of the Spanish Institute of Oceanography). The strain was isolated in 2002 in Girona (Spain), and a single cell was used to establish a monoclonal culture in 2012, yielding clone H7. Culture conditions were an incubation temperature of 19.5 °C ±1 °C with an approximate illumination of 90 µmol·photons·m-2 s-1 and a photoperiod of 12:12 h light:dark (L:D). The cells were grown and maintained in L1 medium (Guillard and Hargraves 1993)
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without added silica. The medium was prepared with Atlantic seawater and adjusted to a salinity of 32 by the addition of sterile distilled water. Synchronization and sampling: When A. minutum clonal culture H7 reached a concentration of 8,000–10,000 cells·mL-1 (corresponding to the exponential growth phase), it
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was transferred to complete darkness for 66 h (synchronization period), following the method of Taroncher-Oldenburg et al. (1997) and Figueroa et al. 2007. Light conditions were then
restored and the cells were inoculated into fresh L1 medium to a concentration of
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approximately 2000 cells mL-1 (1.7 L in total). Sampling started 27 h after inoculation to allow cell recovery and the initiation of division. Supplementary Material Figure S1 summarizes the
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experimental setup and sampling protocol. For flow cytometric analyses, 50-ml samples were taken every hour for 55 h (starting at 11:00 of the first light cycle and ending at 15:00 pm of
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the third light cycle) using an automatic water sampler (AWS, EnviroTech Instruments LLC, Chesapeake, VA, USA), allowing 1 min of programmed soft magnetic shaking just before sampling. In the following, “synchronization” is defined as the period of 66 h of darkness. For
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the determination of absolute cell numbers (cell concentration analyses), two 2-mL samples, the first at 11:00 and the second at 15:00, were collected manually on days 1, 2, 3, 4, 7, 8, and
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9 post-synchronization and fixed with Lugol.
Flow cytometry analyses: Imaging flow cytometry (IFC): Each 50-mL sample was fixed
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with 0.5% formaldehyde for at least 10 min, filtered through a 5.0-µm-pore-size membrane filter (Millipore, Ireland), the fluid discarded and the retained cells resuspended and
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centrifuged at 5000 × g for 5 min. The pellet was resuspended in 2–5 mL of cold methanol and stored for at least 12 h at 4°C to facilitate chlorophyll extraction. The cells were then washed in PBS (pH 7, Sigma-Aldrich, St. Louis, MO, USA) and the pellet was resuspended in staining solution (30 µg·propidium iodide (Sigma-Aldrich) mL-1 and 100 mg· RNaseA (Sigma-Aldrich) mL1
in PBS) for at least 3 h in darkness before analysis. A Flow Sight image flow cytometer (Amnis,
Seattle, WA, USA), with two lasers, emitting at 488 and 405 nm, was used. The samples were run at low speed and data were acquired until 15,000–50,000 events were recorded. Ideas 6.0 (Amnis) and FlowJo 7.6 (Tree Star, Ashland, OR, USA) were used to compute peak numbers, coefficients of variation (CVs), and peak ratios for the DNA fluorescence distributions in a population. CVs were usually below 7; those above 10 were discarded from the analyses. Cell-cycle analyses: The Gradient RMS feature, which enumerates changes of pixel values in the image to measure the focus quality of an image, was used to select focused cells, which have a higher gradient than unfocused cells (Marangon et al. 2012). Aggregates were eliminated using cellular area and nuclear fluorescence and the results verified visually by
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studying the images of both discarded and retained populations. Each cell-cycle phase was delimited by means of a histogram of propidium iodide fluorescence using 405-nm laser excitation, since emission at 488 nm (also recorded) was saturated even at minimum power. The precision of the phase adjustment was compared with that obtained using a Sony iCyt EC800 conventional flow cytometer and a replicated synchronized clonal sample as control.
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The percentages and CVs were similar for the two platforms. Cells were classified as
"individual," in "early cytokinesis," or in "two-cell chains" according to the nuclear-aspect ratio
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(width vs. height of the mask used to more precisely adjust the area to the U-shaped nucleus) and the intensity of nuclear fluorescence. The precision of the adjustment was made manually
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in the control sample by studying the acquired images. With these analyses a general template was built for all samples. The beginning of S phase was set as the time at which its value was half of the maximum, whereas the time of M-phase was estimated as the midpoint of the
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decrease in the percentage of G2/M cells (Dagenais-Bellefeuille et al. 2008). A specific area of an image can be obtained for each feature, which was adjusted in the case of the nuclear area
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by means of a mask to include only well-defined nuclei (IDEAS 6.0, Amnis). Cytoplasmic-tonuclear ratios (CNRs) were calculated as CNR=
Microscopy: Cells in selected samples were fixed for flow cytometry analyses and
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stained with both DAPI (2 µg·mL-1) and Calcofluor to stain the nucleus and thecal plates,
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respectively. In addition, some samples were treated with Liquinox before fixation in ethanol:acetic 3:1 (v/v), The cells were then squashed in a drop of 45% acetic acid onto clean
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microscope slides. The slides were frozen to remove the cover and air-dried before DAPI staining. The cells were examined using a fluorescence microscope (Leica DMR, Germany) and imaged at 1000× magnification using a microscope camera (Axiocam HRC, Zeiss, Germany). Samples for cell-concentration determinations were analyzed in duplicates in an
inverted microscope at 100× magnification using a Sedgwick-rafter chamber. At least 400 cells were counted.
Statistical analyses: Analyses were performed using the statistical and programming
software R 2.1.12 (R Development Core Team, 2012), packages “ggplot2” and “scales”, available through the CRAN repository (www.r-project.org/).
Acknowledgements
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The present work was funded by a FORMAS (Sweden) project to R. I. Figueroa (Formas 215-
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2010-824).
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European)(Dinophyceae) during a summer bloom in Cork Harbour, Ireland (2006). Deep Sea Res Pt II 57:268-278
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Figure Legends
Figure 1. DNA intensity histograms obtained every 2 h for the studied light:dark periods. The dark bar on the left of the panels indicates the dark period. TAS= Time after synchronization.
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Figure 2. Percentage of cells in the different phases of the cell cycle as shown by a trend line and the standard error (gray-shaded area). (A) 27–54 h post-synchronization, (B) 55–79 h postsynchronization, (C) All samples are organized by DNA content level. The dark bars along the bottom indicate the dark period. >2C indicates cells with DNA content higher than 2C.
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Figure 3. Data on cell and nuclear area. (Box-and-whisker plots summarizing cell (A) and nuclear area (B), with the median, first and third quartiles, and minimum and maximum values.
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(C) The regression lines per cell phase.
Figure 4. (A) Cytoplasmic-to-nuclear ratio (CNR) according to cell-cycle phase. The median, first
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and third quartiles, and minimum and maximum values are shown. (B) The CNR of 2C cells and the percentage of 2C cells in early cytokinesis relative to the total population of 2C cells using
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locally weighted regression.
Figure 5. IFC images of cell and nuclear morphologies of cells with DNA contents matching C and S values as seen using bright field (BF) microscopy and after blue (488 nm) and violet (405
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nm) laser excitation.
Figure 6. (A) Classification of 2C cells based on DNA intensity and nuclear aspect ratio. (B)
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Corresponding images of cells as seen in bright field (BF) microscopy, and after blue (488 nm)
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and violet (405 nm) laser excitation.
Figure 7. Percentage of cells in early cytokinesis (dark green) relative to the maximum
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percentage of 2C cells per cycle (light green). The dark bars along the bottom indicate the dark period.
Figure 8. High-magnification (1000×) images of the mitotic process in DAPI- and Calcofluorstained cells, in accordance with the IFC results. Images D–E correspond to nuclei squashes; the other images were obtained from cell suspensions. Scale bars=10 µm.
Supplementary Data
Figure S1. Experimental setup.
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Figure S2. Linear correlations of cell area (μm2) versus time post-synchronization (h) for each group of cells according to their DNA content (>2C indicates cells with a DNA content higher than 2C).
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S1434-4610(15)00002-4 http://dx.doi.org/doi:10.1016/j.protis.2015.01.001 PROTIS 25472
To appear in: Received date: Revised date: Accepted date:
7-7-2014 2-12-2014 8-1-2015
Please cite this article as: Dapena, C., Bravo, I., Cuadrado, A., Figueroa, R.I.,Nuclear and Cell Morphological Changes during the Cell Cycle and Growth of the Toxic Dinoflagellate Alexandrium minutumCell Cycle Analysis of Axeandrium minutum–>, Protist (2015), http://dx.doi.org/10.1016/j.protis.2015.01.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
ORIGINAL PAPER
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Nuclear and Cell Morphological Changes during the Cell Cycle and Growth of the Toxic Dinoflagellate Alexandrium minutum
Instituto Español de Oceanografía (IEO), Subida a Radio Faro 50, 36390 Vigo, Spain
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a
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Carlos Dapenaa, Isabel Bravoa, Angeles Cuadradob; and Rosa Isabel Figueroaa,c,1
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Aquatic Ecology, Biology Building, Lund University, 22362 Lund, Sweden
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Universidad de Alcalá (UAH), Dpto de Biomedicina y Biotecnologia, 28801 Alcalá de Henares, Spain
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Submitted July 7, 2014; Accepted January 8, 2015
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Monitoring Editor: Marina Montresor
Running title: Cell Cycle Analysis of Axeandrium minutum
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Corresponding author; e-mail [email protected]/ [email protected]. (R. I. Figueroa).
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Elucidation of the cell cycle of dinoflagellates is essential to understand the processes leading to their massive proliferations, known as harmful algal blooms. In this study, we used imaging flow cytometry (IFC) to monitor the changes in DNA content and nuclear and cell morphology that occur during clonal growth of the toxic species Alexandrium minutum Halim. Our results indicate that the population was in S phase (C→2C DNA content) during
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the light period, whereas haploid cells with a C DNA content peaked only during a short interval of the dark period. The timing of the phases, identified based on the nuclear
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morphology and cytoplasmic-to-nuclear (CNR) ratio of the cells, suggests that the length of the G2/M phase is regulated by nutrient levels whereas the beginning of S phase is clock
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controlled. In addition we found that up to 7% of individual cells achieved a DNA content higher than 2C, indicative of either zygote formation and replication (homothallism), or of double-haploid cells able to divide (polyploid forms). Cells belonging to different cell cycle
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phases (G1-S-G2/M) could be readily discriminated based on nuclear size. Our study provides evidence of cell-cycle plasticity during clonal growth and unambiguously characterizes the
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cell-cycle phases of this dinoflagellate species.
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Introduction
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Key words: Cell-cycle; cytoplasmic-to-nuclear ratio (CNR); dinoflagellates; growth; nuclear size.
During the cell cycle, eukaryotic cells undergo asexual division to yield two genetically identical daughter cells. The cell cycle proceeds in three stages: i) interphase, marked by three distinct phases: G1, in which growing cells exhibit high metabolic activity; S, during which cellular DNA is replicated; and G2, a temporal gap between S phase and the following division in which cells accumulate the nutrients needed for the division process; ii) mitosis (M phase), during which there is nuclear division, or karyokinesis; and iii) cytokinesis, when cytoplasmic division marks the completion of cell division and thus of the cell cycle. The molecular events that determine a cell's progress through the cell cycle are ordered and directional, such that the cycle is irreversible. There is also an additional stage, called G0, composed of cells that have exited the cycle and stopped dividing. These quiescent cells in Go cannot be differentiated from those in G1, either morphologically or based on their DNA content. However, G0 cells can be activated to re-enter the cell cycle at G1 (see, for example, the review by Pardee 1989). The transition 2
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between different phases of the cell cycle is controlled by cyclins and cyclin-dependent kinases (CDKs), which are highly conserved among eukaryotes. In concert with other species-specific components, cyclins and CDKs regulate the cell cycle itself. Aside from the recognition of the various phases of the cell cycle, eukaryotic diversity in cell-cycle control remains largely unexplored, in part because most of the model species have been unikonts (Harashima et al.
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2013).
The cell cycle is composed of two cycles: the DNA division cycle and the growth cycle,
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which includes the synthesis of all other macromolecules (Vaulot et al. 1986). The DNA
synthesis pattern of many phytoplankton species suggests that a circadian clock controls the
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timing of cell division, with light- and dark-sensitive periods (Chisholm 1981, Chisholm and Brand 1981). Dinoflagellates are unicellular protists, with many species forming harmful algal
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blooms (e.g., Hallegraeff 1993; Smayda 1997). Discrimination between dinoflagellate life- and cell-cycle stages is essential to understand the development of dinoflagellate blooms. During dinoflagellate division, cytokinesis occurs during the day, and karyokinesis/mitosis generally at
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night (Hastings and Sweeney 1964). However, Karentz (1983) observations suggest that there is a huge diversity in the group, different dinoflagellate species differing in their division patterns. For example, DNA synthesis and cell division may be in phase or not, being
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hypothesized that they are in phase in populations with division rates of less than one per day,
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whereas in the others (e.g. Gymnodinium nelsoni or Pyrocystis fusiformis), there is segregation of DNA replication and cell division between different L:D cycles. The same study points to that
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G1, G2 and S length could be specifically regulated among species. Supporting this diversity, DNA synthesis and cell division have been described in dinoflagellates either as a continuous process, similar to that in prokaryotes, or as following a typical eukaryotic G1-S-G2-M phasetransition model (Allen et al. 1975; Polikarpov and Tokareva 1970; Spector et al. 1981), albeit with several singular features (Bhaud et al. 2000). The dinoflagellate nucleus, referred to as the dinokaryon (Rizzo 1991), has many unusual characteristics, most notably its very large genome size. Indeed, the dinoflagellate genome is among the largest of all eukaryotic genomes (Hou and Lin 2009). Dinoflagellate chromosomes, organized by means of a cholesteric liquid crystal structure, are visible throughout the cell cycle (Chow et al. 2010, Rill et al. 1989), but in many respects their morphology resembles that seen in other eukaryotes (Figueroa et al. 2014). The chromosomes divide via closed mitosis, with a permanent nuclear envelope and through cytoplasmic channels (Soyer-Gobillard et al. 1999), but they lack detectable histones, despite the fact that recent studies demonstrated the existence of all core histone mRNA sequences (Lin et al. 2010, Roy and Morse 2012). The characteristics of the dinoflagellate nucleus have
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been attributed to the acquisition of prokaryotic genes during evolution, which may also explain the diversity and relatively slow growth rates of dinoflagellates. The latter has been attributed to their having a Rubisco enzyme, which inefficiently distinguishes CO2 from O2 and probably originated from anaerobic protobacteria (Wong and Kwok 2005).
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In the dinoflagellate Lingulodinium polyedrum, S phase, and not mitosis, was gated by the circadian clock (Homma and Hastings 1989; Lidie et al. 2005). Nonetheless, the results of Dagenais-Bellefeuille et al. (2008) in this species are consistent with a circadian clock
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controlling both the G2/M and S phases. Nutrients have been considered as potential modulators of the cell cycle phases, but the length of the G2 phase in phytoplankton growing
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under different nutritional levels remained constant in the study by Olson and Chisholm (1986).
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Blooms of the cosmopolitan and toxin-producing dinoflagellate Alexandrium minutum Halim occur worldwide in shallow systems such as estuaries, bays, lagoons, and harbors (Bravo
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et al. 2010; Garcés et al. 2004; Giacobbe et al. 1996; Touzet et al. 2010). Their negative effects, including production of the toxin that causes paralytic shellfish poisoning, have been welldocumented in humans and in other species. Growth in Alexandrium, including A. minutum, is
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mainly achieved through asexual division (mitosis). Sexual reproduction can be induced by,
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among other factors, phosphate deficiency and apparently involves more than two compatible mating types (complex heterothallism), given that self-fertilization in clonal strains has yet to
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be observed (Figueroa et al. 2007). In dinoflagellate ecology, sexual reproduction has several benefits in addition to the potential for genetic recombination; mainly in those species in which leads to resting cyst formation (see Bravo and Figueroa 2014 and references therein). Resting cysts are benthic forms that are highly resistant to adverse conditions, and they can remain viable in the sediments for up to 100 years (Miyazono et al. 2012). Germination and subsequent divisions (meiosis) of the resting cyst restore its haploid life-stage. Apart from the cyst stage, which in most dinoflagellate species is morphologically very different from planktonic forms, the life- and cell-cycle stages of Alexandrium are in general very similar. Resting cysts are well characterized by the absence of flagella, a double cell wall, and the presence of brownish pigment deposits. However, most characteristics considered useful in distinguishing flagellated stages (vegetative stages, gametes, and mobile zygotes) are in fact misleading. For example, regarding size, gametes are generally considered to be smaller, and zygotes larger than vegetative cells (see the revision by Bravo and Figueroa 2014). However, for A. minutum, gametes with a size similar to that of the population average and smaller than zygotes have been observed (Figueroa et al. 2007). Immediately after their formation, mobile 4
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zygotes possess two trailing flagella, unlike the uniflagellate cells of vegetative stages. However, this characteristic is easily lost by cells under stress or after cell fixation. Therefore, the identification of Alexandrium life and cell cycle stages is often very challenging but essential to understand growth and survival strategies of these organisms in their natural
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habitats. In their study of toxin production during the cell cycle in Alexandrium fundyense, Taroncher-Oldenburg et al. (1997) showed that prolonged darkness arrested the cells in a
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light-dependent part of the cell cycle (G0), which prevented both progression through G1 and division. DNA synthesis (S phase) started just before the beginning of the dark period, reaching
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a maximum that coincided with the middle of G2/M phase. Earlier studies showed that at the G1/S boundary, eukaryotic cells must pass a size threshold, whereas at the G2/M boundary a
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prerequisite for further cell-cycle progression is the completion of DNA synthesis (Elledge 1996). In this study, we asked whether this pattern was also relevant for A. minutum and if cells of this species could be unambiguously characterized according to their cell-cycle-specific
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morphology and their nuclear and cellular sizes. To answer this question and to determine the timing of each cell-cycle phase, we used imaging flow cytometry (IFC) and followed nuclear and cellular morphologies to study a darkness-synchronized clone of the dinoflagellate
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Alexandrium minutum over a period of 53 h.
Results
Growth
During the first light:dark period (28–48 TPS or h post-synchronization) the number of cells doubled, from 3290 ± 42.4 cells·mL-1 to 6313 ± 245.4 cells·mL-1 (0.94 div·day-1). During the second light:dark period, growth was slower, with the population increasing from 6313 ± 245.4 cells·mL-1 to 8735 ± 49.5 cells·mL-1 (0.47 div·day-1). Although the cell cycle was evaluated by flow cytometry for only the first 53 h, growth of the clonal culture was followed until day 8 post-synchronization, reaching over 30,000 cells·mL-1. DNA Content The cells were classified according to their DNA content, as determined by the peaks in the nuclear fluorescence histogram of Figure 1, as C, C→2C, and 2C. At the beginning of the study 5
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(28 h TPS) (Fig. 1A) and under light conditions (08:00–20:00), these histograms showed the presence of one main peak, with a relatively high coefficient of variation (CV), which widened over time. However, as the cells progressed through the dark period (20:00), two additional peaks clearly appeared. The one to the left of the first peak corresponded to G1 (1C DNA content) and the other, at the far right, to 2C (2C/C=1.95). Consequently, the main peak
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observed under light conditions was that of S phase (C→2C). S ll during the dark period, at
04:00, or 44 h TPS in Fig. 1, the G1 peak shifted slightly, towards the position of the S-phase
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peak. The latter was maintained in the second light period, starting at 48 h TPS (Fig. 1), until the dark period (20:00 or 60h TPS), during which time the size of the 2C peak started to
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increase. Note that the main position of the cell population during the second light period (C→ 2C peak) was closer to the 1C position than during the first light period. Additionally, peaks corresponding to 1C and 2C occurred later, 2 h after the onset of the dark period (22:00),
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whereas the decrease in the 2C peak stopped 2 h after the beginning of the third light period, rather than during the dark period, as observed before. The size of the 2C peak was
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significantly smaller in the second than in the first dark period. At the beginning of the third light period, the main peak shifted slightly towards the C→2C peak, as observed before; however the displacement was again less than during the first light period.
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Consistent with the different peak sizes shown in Figure 1, the maximum percentage of
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cells in G1 or S phase or undergoing division differed substantially between the light:dark periods studied (Fig. 2A, B), although the overall tendencies were similar (Fig. 2C): Maximums
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of 2C cells and cells with a >2C DNA content and minimums of C→2C cells were detected during darkness, whereas C→2C cells reached their maximum numbers during the light period. In the first light:dark period, the percentages of cells in S and G2/M were markedly higher than during the following period. This finding was supported by the fact that the best curve fit was obtained with separate plots, i.e., 27–54 h TPS (Fig. 2A) and 55–79 h TPS (Fig. 2B). From the graphs in Figure 2, we estimated that the beginning of S phase was at 07:00 in both cycles. G2/M phase started at 18:00 and lasted until 06:00 in the first cycle but until 09:00 in the second. S phase was also longer (~2 h) in the second cycle. Individual cells with a DNA content >2C were detected in both cycles and their pattern of appearance in the dark period was similar to that of 2C cells, but with a much smaller percentage of cells: maximally 7.7% in the first cycle and 7.5% in the second cycle. Morphology
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Cell and nuclear area. Figure 3 shows the average (N>10,000) cellular (Fig. 3A) and nuclear (Fig. 3B) areas together with the cell cycle data for individual cells. The values corresponding to the different phases did not overlap, with the largest differences between phases recorded at the nuclear level. Cell and nuclear areas were strongly correlated, following a linear relationship in all cell-cycle phases and when data from these phases were pooled (Fig.
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3C). After the peak in 2C cells, the average cell size after division (C) decreased until around
400 μm2 in the first light:dark period but reached close to 300 μm2 in the second (Fig. 3A), in
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which fewer cells underwent division (Fig. 2A-B). This fact is shown by a negative correlation in the size of cells over time, observed for all DNA contents unless in cells>2C (Supplementary
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Material Fig. S2). The cytoplasmic-to-nuclear ratio (CNR) changed over time and according to DNA content (Fig. 4A), trending to a linear correlation (R2=0.514). The CNRs of cells with C DNA content overlapped slightly with those of cells with C→2C, even if the median values differed
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(Fig. 4A). Pairwise comparisons using t tests showed highly significant differences in CNR values between groups with different DNA content (Supplementary Material Table S1)
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Nuclear morphology during the different cell-cycle stages. The nuclei of cells analyzed by IFC showed either a roundish or a very weakly lobulated morphology for those with a C DNA content, but a more typical U-shaped morphology characteristic of the Alexandrium genus for
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those with a C→2C DNA content, indicative of S phase (Fig. 5). Three groups were defined in a
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specific study of 2C cells: “individual,” “in early cytokinesis,” and “in two-cell chains” (Fig. 6A). The nuclear morphologies were consistent with the cellular classification (Fig. 6B), as individual
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cells had 2C either U-shaped or roundish nuclei; cells in early cytokinesis had either two C nuclei or one elongated 2C nucleus; and two-cell chains consisted of two differentiated cells with two physically separated C nuclei. The percentage of individual cells in division was seen as a distinct peak that corresponded to the maximum of 2C cells per cycle (Fig. 7). The IFCbased determination of cells in early cytokinesis indicated the onset of mitosis at a similar time than the maximum value of CNR of 2C cells, which peaked at cell division (Fig. 4B). According to the CNRs and the curve followed by cells in early cytokinesis, mitosis occurred at 21:00– 23:00 (37-39 h post-synchronization in the first cycle and at 22:00–01:00 (62–65 h postsynchronization in the second cycle (Figs 4B and 7). Using more detailed images, taken at higher magnification, together with those obtained by IFC, we reconstructed the cell cycle process. Figure 8A, B shows cells smaller than the minimum size needed to enter S phase. The nuclei of these cells are small and U-shaped, with very shallow concavities (Fig. 8A), and are roundish in appearance depending on the angle of view (Fig. 8B). A 2C stage, with a well-developed U-shaped nucleus, is shown in Figure 8C; a 7
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putative metaphase cell, with well-separated chromosomes, is shown in Figure 8D, and a late metaphase cell, with double-chromatid chromosomes (arrows), in Figure 8E. Our reconstruction of anaphase and telophase is provided in Figure 8F–H, which shows these two processes until the formation of a two-cell chain, a mitotic product that is subsequently broken to restore the individuality of the cells. The cell depicted in Figure 8J has a round nucleus and
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decondensed chromosomes, similar to many of the 2C individual cells seen in Figure 6B. Lastly, based on its size and that of its nucleus, the cell in Figure 8K is probably one with a DNA
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content >2C.
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Discussion
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The study of dinoflagellate proliferation has considerable interest because it can lead to the formation of Harmful Algal Blooms (HABs), with adverse effects on public health, seafood safety and aquaculture (Smayda 2002). The rate of division of a given population of cells is
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under tight control of cell cycle, a key element of this study. The present work focuses in the
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cell cycle characterization of the cosmopolitan PST-producing species Alexandrium mintum, in order to determine both the timing of the DNA content changes and how cell growth,
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estimated by cell and nuclear areas, relate to the progression and stages of the cell cycle. DNA Content Changes
Many phytoplankton species show circadian regulation of cell cycle (Chisholm and Brand 1981; Homma and Hastings 1989; Sweeney and Hastings 1958), which exhibit three main characteristics: they are endogenously generated, self-sustaining, and persist under constant conditions, with light vs. dark conditions and temperature among the most well-studied (see the review of McClung 2006 and references therein). The “escape from light hypothesis” (Pittendrigh 1993) posits that organisms gain an advantage by carrying out light-sensitive processes, such as DNA replication, during the dark portion of the diel cycle. However, in some dinoflagellates, such as Lingulodinium polyedrum (Dagenais-Bellefeuille et al. 2008; Homma and Hastings 1989), S phase occurs during the dark period, whereas in other species, such as Amphidinium operculatum (Leighfield and Van Dolah 2001; Van Dolah and Leighfield 1999), Karenia brevis (Van Dolah et al. 2007), and Alexandrium fundyense (Taroncher-Oldenburg et al. 1997) it takes place, at least in part, during the day. The last authors reported that immediately
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after the cells are released from prolonged darkness, they enter a nonproliferation period equivalent to one generation time, as result of arrest in G0. These cells are too small to enter and progress through G1. For that reason, our first sampling was 27 h after darkness release and it showed that in A. minutum S-phase occurs entirely during the light period. Moreover, in the second light:dark period of our study, when only a fraction of the cells in the population
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divided, the average DNA content of the population still occurred within the S phase (C→2C),
during the light period. A peak corresponding to cells with a C DNA content was observed only
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after G2/M, under conditions of darkness. Interference due to RNA content can be ruled out in these observations given the high concentrations of RNAase used in the study to specifically
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avoid this problem. Interestingly, and in contrast to L. polyedrum (Dagenais-Bellefeuille et al. 2008), the timing and length of S and G2 in A. minutum varied with growth, as both phases were shorter in the first cycle, when the growth rate was higher, and longer in the second,
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when there was less growth of the population. DNA replication and mitosis are controlled by specific proteins whose rate of synthesis changes during the cell cycle (Evans et al. 1983; Giard
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et al. 1973), and growth-dependent variations in the durations of S and G2 have been demonstrated in different cell lines, e.g., in yeasts. In that study, Slater el al. (1977) examined the relation between the lengths of the cell-cycle phases and the growth rate of the culture.
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The largest variability was in the length of G1 (10-fold vs. nearly 2-fold in both S and G2). In
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dinoflagellate species with division rates of less than one per day , the cell cycle is said to be ruled by the circadian clock, meaning that there is a temporal window through which cells that
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have reached a minimum size threshold can proceed through the cycle (Elledge 1996). This implies cell-cycle control at the S and G2 phases. Indeed, in our experiment these two phases similarly varied between light:dark periods, as the difference in the duration of either one was approximately 2 h. Cell growth has to be coordinated with the cell division cycle to allow an appropriate response to nutrient availability (Wong and Kwok et al. 2005). Our results show that all cells duplicated during the first light:dark period but the population decreased their growth rate during the following one, indicating a progressive loss of cell cycle synchrony. The maximum division rate achieved in the first light:dark period (A. minutum usually grows at lower rates) is probably not possible to maintain over long periods of time, because metabolic needs and nutrient uptake become imbalanced when synchronization is lost (e.g., TaroncherOldenburg et al. 1997, Vaulot et al. 1996). Supporting this theory, during the 53 h of our study the trend was a decrease in the average area of C cells, which suggests nutritional depletion. Thus, dinoflagellates are apparently able to rapidly adapt to the available nutritional resources, modulating their cell-cycle progression by increasing the length of both G2 and S in response
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to the number of cells in G1 that are able to reach the minimum threshold size needed for entry into S phase. Another implication of our results is related to the validity of the estimations of cellular DNA content in some dinoflagellate species. Those estimates were mainly based on
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unsynchronized cultures sampled during light conditions (e.g., LaJeunesse et al.2005). However, according to the results of our study, the DNA content of some dinoflagellate species may be overestimated by this method when S phase starts immediately before the beginning
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of the light period.
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Additionally, another novel result in the present work was the detection of individual cells with a >2C DNA content. One explanation could be related to the sexual cycle of this species. A. minutum has been described as a heterothallic species (Figueroa et al. 2007), as
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two different clones were shown to be necessary to produce sexual resting cysts. However, that study was unable to determine why some zygotes encyst while others divide. We think
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that the existence of cells with a >2C DNA content in the clonal culture (in which no resting cysts were ever observed) is best explained by low levels of homothallism and zygote division (by mitosis or meiosis), as suggested by the fact that cells with a DNA content >2C show a
d
similar trend of 2C cells, with peaks during the dark period. Thus, our study is the first to show
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a circadian rhythm for the division of dinoflagellate cells with a ploidy level different than that of haploid cells. While an alternative explanation is the existence of double haploid cells able
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to divide, as a consequence of a possible polyploidization event, we think that the first hypothesis is more plausible because the percentage of these cells increases dramatically following a sexual cross, suggesting that sexuality and zygote formation underlie the origin of cells forming this population (Figueroa et al., unpubl. observ.). Cell and Nuclear Size during the Cell Cycle Nuclear size and shape are carefully controlled in eukaryotic cells, with aberrations in either leading to altered cellular states such as cancer (see the review by Huber and Gerace 2007 and references therein). Cell division requires an increase in the cell surface area:volume ratio (Oney et al. 2005); indeed, there is a well-established correlation between cell and nuclear size, expressed as the CNR, with changes in DNA content altering cell volume (see, for instance, the work in yeasts by Jorgensen et al. 2007). Early observations formed the basis of the “karyoplasmic” ratio theory, according to which eukaryotic cells actively maintain a fixed ratio of nuclear to cell volume (Cavalier-Smith 2005; Wilson 1925). The validity of this hypothesis has been confirmed in cells with a broad range of genetic backgrounds and placed in a wide
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variety of growth conditions, including those that alter cell volume and DNA content. But to our knowledge ours is the first study to confirm this relationship in dinoflagellates, i.e., the eukaryotes with the highest DNA content (Lin 2011). The CNR is an important indicator of cell stability. For example, for the multifunctional
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protein survivin, an inhibitor of apoptosis and a breast cancer biomarker, the CNR was shown to be an important predictor of outcome in tamoxifen-treated patients (Rexhepaj et al. 2010). The variations in the CNRs in our study were mainly but, as evidenced by the data dispersion,
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not exclusively limited to DNA content, consistent with reports of factors besides the amount of DNA in the nucleus affecting the CNR. For example, nuclear size might also be subject to
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regulation depending on cell-type-specific chromatin condensation (Swanson et al. 1991). The CNRs of A. minutum cells in G1 and S overlapped but increased significantly in cells with a DNA
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content ≥2C.
The morphological discrimination between dinoflagellate life- and cell-cycle stages has
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been difficult if not impossible but it is essential if we are to understand dinoflagellate blooming patterns in the field. Our results show that cell and nuclear area, as well as the CNR, can be used to efficiently distinguish between the cell-cycle stages of the dinoflagellate A.
d
minutum. Further studies are necessary to determine whether these relationships are
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maintained between genotypes, along the growth curve, and under different environmental conditions, and to design parameters able to accurately evaluate the growth of a bloom and
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forecast its development potential.
Methods
Cell proliferation conditions: The dinoflagellate strain used in this study was A. minutum strain VGO577, regularly maintained at the Centro Oceanográfico de Vigo (CCVIEO; Culture Collection of Harmful Microalgae of the Spanish Institute of Oceanography). The strain was isolated in 2002 in Girona (Spain), and a single cell was used to establish a monoclonal culture in 2012, yielding clone H7. Culture conditions were an incubation temperature of 19.5 °C ±1 °C with an approximate illumination of 90 µmol·photons·m-2 s-1 and a photoperiod of 12:12 h light:dark (L:D). The cells were grown and maintained in L1 medium (Guillard and Hargraves 1993)
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without added silica. The medium was prepared with Atlantic seawater and adjusted to a salinity of 32 by the addition of sterile distilled water. Synchronization and sampling: When A. minutum clonal culture H7 reached a concentration of 8,000–10,000 cells·mL-1 (corresponding to the exponential growth phase), it
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was transferred to complete darkness for 66 h (synchronization period), following the method of Taroncher-Oldenburg et al. (1997) and Figueroa et al. 2007. Light conditions were then
restored and the cells were inoculated into fresh L1 medium to a concentration of
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approximately 2000 cells mL-1 (1.7 L in total). Sampling started 27 h after inoculation to allow cell recovery and the initiation of division. Supplementary Material Figure S1 summarizes the
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experimental setup and sampling protocol. For flow cytometric analyses, 50-ml samples were taken every hour for 55 h (starting at 11:00 of the first light cycle and ending at 15:00 pm of
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the third light cycle) using an automatic water sampler (AWS, EnviroTech Instruments LLC, Chesapeake, VA, USA), allowing 1 min of programmed soft magnetic shaking just before sampling. In the following, “synchronization” is defined as the period of 66 h of darkness. For
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the determination of absolute cell numbers (cell concentration analyses), two 2-mL samples, the first at 11:00 and the second at 15:00, were collected manually on days 1, 2, 3, 4, 7, 8, and
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9 post-synchronization and fixed with Lugol.
Flow cytometry analyses: Imaging flow cytometry (IFC): Each 50-mL sample was fixed
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with 0.5% formaldehyde for at least 10 min, filtered through a 5.0-µm-pore-size membrane filter (Millipore, Ireland), the fluid discarded and the retained cells resuspended and
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centrifuged at 5000 × g for 5 min. The pellet was resuspended in 2–5 mL of cold methanol and stored for at least 12 h at 4°C to facilitate chlorophyll extraction. The cells were then washed in PBS (pH 7, Sigma-Aldrich, St. Louis, MO, USA) and the pellet was resuspended in staining solution (30 µg·propidium iodide (Sigma-Aldrich) mL-1 and 100 mg· RNaseA (Sigma-Aldrich) mL1
in PBS) for at least 3 h in darkness before analysis. A Flow Sight image flow cytometer (Amnis,
Seattle, WA, USA), with two lasers, emitting at 488 and 405 nm, was used. The samples were run at low speed and data were acquired until 15,000–50,000 events were recorded. Ideas 6.0 (Amnis) and FlowJo 7.6 (Tree Star, Ashland, OR, USA) were used to compute peak numbers, coefficients of variation (CVs), and peak ratios for the DNA fluorescence distributions in a population. CVs were usually below 7; those above 10 were discarded from the analyses. Cell-cycle analyses: The Gradient RMS feature, which enumerates changes of pixel values in the image to measure the focus quality of an image, was used to select focused cells, which have a higher gradient than unfocused cells (Marangon et al. 2012). Aggregates were eliminated using cellular area and nuclear fluorescence and the results verified visually by
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studying the images of both discarded and retained populations. Each cell-cycle phase was delimited by means of a histogram of propidium iodide fluorescence using 405-nm laser excitation, since emission at 488 nm (also recorded) was saturated even at minimum power. The precision of the phase adjustment was compared with that obtained using a Sony iCyt EC800 conventional flow cytometer and a replicated synchronized clonal sample as control.
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The percentages and CVs were similar for the two platforms. Cells were classified as
"individual," in "early cytokinesis," or in "two-cell chains" according to the nuclear-aspect ratio
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(width vs. height of the mask used to more precisely adjust the area to the U-shaped nucleus) and the intensity of nuclear fluorescence. The precision of the adjustment was made manually
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in the control sample by studying the acquired images. With these analyses a general template was built for all samples. The beginning of S phase was set as the time at which its value was half of the maximum, whereas the time of M-phase was estimated as the midpoint of the
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decrease in the percentage of G2/M cells (Dagenais-Bellefeuille et al. 2008). A specific area of an image can be obtained for each feature, which was adjusted in the case of the nuclear area
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by means of a mask to include only well-defined nuclei (IDEAS 6.0, Amnis). Cytoplasmic-tonuclear ratios (CNRs) were calculated as CNR=
Microscopy: Cells in selected samples were fixed for flow cytometry analyses and
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stained with both DAPI (2 µg·mL-1) and Calcofluor to stain the nucleus and thecal plates,
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respectively. In addition, some samples were treated with Liquinox before fixation in ethanol:acetic 3:1 (v/v), The cells were then squashed in a drop of 45% acetic acid onto clean
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microscope slides. The slides were frozen to remove the cover and air-dried before DAPI staining. The cells were examined using a fluorescence microscope (Leica DMR, Germany) and imaged at 1000× magnification using a microscope camera (Axiocam HRC, Zeiss, Germany). Samples for cell-concentration determinations were analyzed in duplicates in an
inverted microscope at 100× magnification using a Sedgwick-rafter chamber. At least 400 cells were counted.
Statistical analyses: Analyses were performed using the statistical and programming
software R 2.1.12 (R Development Core Team, 2012), packages “ggplot2” and “scales”, available through the CRAN repository (www.r-project.org/).
Acknowledgements
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The present work was funded by a FORMAS (Sweden) project to R. I. Figueroa (Formas 215-
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2010-824).
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Figure Legends
Figure 1. DNA intensity histograms obtained every 2 h for the studied light:dark periods. The dark bar on the left of the panels indicates the dark period. TAS= Time after synchronization.
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Figure 2. Percentage of cells in the different phases of the cell cycle as shown by a trend line and the standard error (gray-shaded area). (A) 27–54 h post-synchronization, (B) 55–79 h postsynchronization, (C) All samples are organized by DNA content level. The dark bars along the bottom indicate the dark period. >2C indicates cells with DNA content higher than 2C.
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Figure 3. Data on cell and nuclear area. (Box-and-whisker plots summarizing cell (A) and nuclear area (B), with the median, first and third quartiles, and minimum and maximum values.
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(C) The regression lines per cell phase.
Figure 4. (A) Cytoplasmic-to-nuclear ratio (CNR) according to cell-cycle phase. The median, first
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and third quartiles, and minimum and maximum values are shown. (B) The CNR of 2C cells and the percentage of 2C cells in early cytokinesis relative to the total population of 2C cells using
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locally weighted regression.
Figure 5. IFC images of cell and nuclear morphologies of cells with DNA contents matching C and S values as seen using bright field (BF) microscopy and after blue (488 nm) and violet (405
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nm) laser excitation.
Figure 6. (A) Classification of 2C cells based on DNA intensity and nuclear aspect ratio. (B)
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Corresponding images of cells as seen in bright field (BF) microscopy, and after blue (488 nm)
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and violet (405 nm) laser excitation.
Figure 7. Percentage of cells in early cytokinesis (dark green) relative to the maximum
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percentage of 2C cells per cycle (light green). The dark bars along the bottom indicate the dark period.
Figure 8. High-magnification (1000×) images of the mitotic process in DAPI- and Calcofluorstained cells, in accordance with the IFC results. Images D–E correspond to nuclei squashes; the other images were obtained from cell suspensions. Scale bars=10 µm.
Supplementary Data
Figure S1. Experimental setup.
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Figure S2. Linear correlations of cell area (μm2) versus time post-synchronization (h) for each group of cells according to their DNA content (>2C indicates cells with a DNA content higher than 2C).
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