TY - JOUR
AU - Wada,, Masamitsu
AB - Abstract The haploid gametophyte generation of ferns is an excellent experimental material for cell biology studies because of its simple structure and high sensitivity to light. Each step of the developmental process, such as cell growth, cell cycle and the direction of cell division, is controlled, step by step, by light, unlike what happens in complex seed plant tissues. To perform analyses at the cell or organelle level, we have developed special tools, instruments and techniques, such as a cuvette suitable for repeated centrifugation in particular directions, microbeam irradiators for partial cell irradiation and single-cell ligation technique to create enucleated cells. Some of our main discoveries are as follows: (1) changes in the intracellular position of the nucleus in long protonemal cells by centrifugation revealed that the nuclear position or a factor(s) that is/are co-centrifuged with the nucleus is important for the decision regarding the place of the formation of preprophase bands and the timing of their disappearance, which determines the position where the new cell wall attaches to the mother cell wall; (2) even within a single cell, various phenomena could be induced by blue or red light, with the localization of the blue or red light receptors being different depending on the phenomenon; (3) de novo mRNA synthesis is not involved in the signal transduction pathways underlying light-induced chloroplast movements. In this review article, various microscopic techniques, in addition to the results of physiology studies in fern gametophytes, are described. Adiantum capillus-veneris, chloroplast, gametophyte, photomorphogenesis, phototropin Introduction Adaptation of plants to environmental stresses during evolution Land plants originated from aquatic algae. Once they moved on to the land, plants expanded their niches on earth and proliferated by adapting to dry conditions and to strong sunlight, especially to its ultraviolet component [1]. Cell walls, central to determining the shape of plants, and the evolution of anchoring devices such as rhizoids or roots, resulted in the sessile nature of land plants, requiring plants to adapt by responding to environmental factors, such as extremes of temperature, light and water availability [2,3]. Of these factors, light is the most useful one for plant life because of its various components, such as wavelength, intensity and duration of irradiation, direction and vibration plane of the incident light, on/off nature of light, day length and seasonal changes [4–6]. Plants use these light quality and quantity traits not only as energy for photosynthesis but also as sources of information, for detecting changes in the local environmental conditions in order to respond by altering their behavior. Light is a researcher-friendly experimental tool by which to control, step by step, the elemental processes of developmental and physiological phenomena in plants [5–7]. Distinct from chemicals or plant hormones, light can be applied non-destructively and instantaneously from outside without any residual effects when the light source is switched off. Light activation can be restricted to the area where photoreceptors exist. When light is used as information to detect environmental condition, short pulse of low intensity light is enough, so that the influence of photosynthetic products is negligible. Light as a factor in controlling photomorphogenesis in plants The germination of spores and seeds, development and differentiation of thalli and seedlings, transition from the vegetative to the reproductive stage, and the cellular and subcellular components of physiological processes such as cellular growth, development of chloroplasts, as well as the underlying differential gene expression, are all controlled, to a greater or lesser extent, by light [4–6,8]. The light-mediated development of plant structure and form is known as photomorphogenesis. Light-dependent phenomena in plants can be readily observed in our daily lives, with plants placed on a window sill showing phototropic responses or leaf spreading towards the window. Photomorphogenesis is mediated by photoreceptors. Red light is absorbed by phytochromes [9,10] while blue light is absorbed by the blue-light receptors phototropins, cryptochromes and/or the ZTL/LKP/FKF family of proteins [11–13], with UVR8 having recently been identified as a UVA receptor [14,15]. Understanding of the basic mechanisms underlying developmental and physiological processes can contribute to resolving vital problems such as food shortages in the near future and today’s environmental pollution, with results from basic as well as applied research contributing to solutions to such pressing issues. Since this review is published in the journal ‘Microscopy,’ not only the experimental results but also the development and use of microscopic techniques in photobiology will be explained in detail, although these techniques are usually not described in reviews. Materials and Methods Plants for experimental material The rate of progress in research frequently depends on the experimental materials we choose to study. In the selection of plant materials on which to perform photomorphogenesis experiments, valuable characteristics include high sensitivity to light, simple organization for ease of irradiation and observation of light effects, high efficiencies of mutant induction and screening of mutant lines, small genome size with minimal gene duplication, and easy knockout and knock-in of genes, although materials that exhibit all of these traits are extremely rare. Even model plants, such as Arabidopsis thaliana, have merits for some types of experiments but also demerits for other purposes. Therefore, we have to select plant material which satisfies most of the characteristics that are essential for our own experiments. A. thaliana, whose genome sequence has been completely sequenced [16], is an excellent material for genetic analyses, but its multi-cellular structure is not appropriate for experiments at the cellular or subcellular levels. The gametophytic generation in ferns, mosses, and liverworts has a simple linear or 2D structure, with a single cell layer, at least during their early developmental stages, which is not surrounded by any tissues, so that light irradiation and observation of the results are straightforward. The gametophytes of the maidenhair fern, Adiantum capillus-veneris, are light sensitive and every elementary process of their development, such as cell elongation, progress of the cell cycle, and direction of the protonemal growth, is strongly controlled by light [7]. The problem undermining the use of fern gametophytes, however, lies in the fact that genetic analysis by stable gene transformation has not yet been established for fern gametophytes, although transient gene transfer by particle bombardment can be achieved [17]. In the common liverwort, Marchantia polymorpha, various techniques of genetic manipulation have been developed [18] and its genome size is very small because of the low genetic redundancy [19]. M. polymorpha is now becoming one of the best model plants [19,20], although few physiological processes can be observed in terms of photomorphogenesis. The spreading earthmoss, Physcomitrella patens, is also used frequently as a model plant [21], although its high level of gene duplication and resulting functional redundancy prevents it from being an ideal model plant [22]. Here, I will review the experimental results obtained in my laboratory using A. capillus-veneris gametophytes. Please refer to a number of review articles highlighted here for more details [7,23–26]. Life cycle of ferns The fern life cycle shows alternation of generation (Fig. 1). Spore germination, thread-like protonema growth (Fig. 1a), heart-shaped prothallium growth (Fig. 1b), and differentiation into antheridia and archegonia (Fig. 1c) occur during the gametophytic generation. After fertilization of an egg cell by a sperm, the sporophyte generation starts (Fig. 1d). The former is the haploid generation while the latter is the diploid generation. The simple structure of the haploid gametophyte phase is valuable material for experimental studies. Fig. 1. View largeDownload slide Life cycle of Adiantum capillus-veneris. a, protonema; b, prothallium; c, antheridium and archegonium; d, young sporophyte growing on gametophyte. Redrawn the figure in [24]. Fig. 1. View largeDownload slide Life cycle of Adiantum capillus-veneris. a, protonema; b, prothallium; c, antheridium and archegonium; d, young sporophyte growing on gametophyte. Redrawn the figure in [24]. Culture of fern gametophytes Culture medium For experimental use, standardization of the gametophyte quality is best-achieved by cultivation on the surface of an agar medium solidified with 0.5% agar. Medium solutions with minerals, such as Murashige and Skoog medium or White medium, are needed, especially for long-term cultivation, but even tap water is sufficient for short-term cultivation. We generally use 1/10 dilution of the Murashige and Skoog basal medium [27]. Spore sterilization Spores have to be sterilized with 1/10 dilution Antiformin (commercially available Sodium hypochlorite solution) containing 0.1% (v/v) Triton X-100. If a large quantity of spores has to be sterilized, e.g. for biochemical studies, the spores are suspended in Antiformin plus Triton X-100 solution in a centrifuge tube and shaken for several minutes before being centrifuged, after which the supernatant is discarded and the spores pellet is washed several times with sterilized distilled water by repeated centrifugation and re-suspension. When only a small number of gametophytes are needed, the spores are spread out on a weighing paper, from which they are picked up with a nichrome wire (either loop or straight wire) sterilized by flaming then cooling by wetting the wire with the Antiformin solution, keep it for 30 s for spore sterilization, then used to inoculate the spores onto the surface of the solid medium (see Fig. 2b of [27]). Fig. 2. View largeDownload slide Schematic illustration of a culture box with a petri dish. The box with a narrow window set with a red acrylic plate at one side was made from a light-tight black acrylic plate. The glass petri dish in the box contains agar medium with protonemal cells covered with narrow cover glasses on the surface of the medium, so that the protonemal cells grow between the agar surface and the cover glass towards the red light from the side window. Fig. 2. View largeDownload slide Schematic illustration of a culture box with a petri dish. The box with a narrow window set with a red acrylic plate at one side was made from a light-tight black acrylic plate. The glass petri dish in the box contains agar medium with protonemal cells covered with narrow cover glasses on the surface of the medium, so that the protonemal cells grow between the agar surface and the cover glass towards the red light from the side window. Cultivation Gametophyte cells (i.e. spores, protonemata and prothallia) growing on an agar surface cannot be observed in detail because of the refraction of light penetrating from the air (outside the cells) to the water (inside the cells). This also means that those cells on the agar surface cannot be illuminated evenly with light. For precise illumination by excitation light and exact observations, the cells should be covered with a narrow (3-mm wide) coverslip and cultivated between the agar surface and the glass not to be exposed in the air. The narrow coverslip is recommended because of the need for aeration in order to supply dissolved air to the gametophytes (we use custom-made coverslips, 3 × 18 mm in size) (Fig. 2). Culture dish and container In the case of irradiation with polarized light, glass dishes are preferable to plastic dishes as the latter disturb the vibration plane of polarized light. We use 3-cm diameter glass petri dishes. The dishes containing spores are placed in a light-impenetrable box made of a black acrylic plate with a side window equipped with a red plastic plate in order to irradiate the spores with red light from a horizontal direction. We use a hand-made small box (length × width × height, 130 × 50 × 30 mm; Fig. 2, see also Fig. 2a of [27]). Methods in photomorphogenesis studies To perform studies on photomorphogenesis, there are several techniques specific for light treatment (refer to [28]). Firstly, all experiments should be carried out in a dark room, the inside of which should be painted black to reduce the influence of any stray light. Only the appropriate parts of the plant material, the light response of which is to be investigated, are irradiated. In this procedure, any light other than the excitation light, such as stray light from a light source, a light path through a microbeam irradiator, or even from a monitor screen, must be blocked. However, the researcher needs to see the materials and tools to carry out the experimental procedures and to observe the experimental results, using a ‘safe light’ that is not influential (or is less influential) to the plant material. The safe light is ideally an infrared light that cannot be absorbed by plant cells, or dim green light at a wavelength of around 550 nm, which plant photoreceptors such as chlorophylls (Fig. 3c), phytochromes (Fig. 3b) and blue-light absorbing pigments like phototropins (Fig. 3c), do not absorb or absorb little, but to which human eyes have the greatest possible sensitivity (Fig. 3a). The dim green safe light can be obtained by white fluorescent light passing through amber plus blue filters [29] but currently a green LED of 550 nm can easily be obtained, although the light intensity needs to be reduced using neutral density filters. Any monitor screens in the experimental area should also be covered with this filter set. To avoid light penetration into a dark room when researchers go in and out of the room during the experiments, the dark room should have a light-tight small front chamber. Consequently, there are some specific methods and techniques to which attention should be paid to perform photomorphogenesis experiments, but not for other research fields. Here, I will summarize fundamental issues for photomorphogenesis studies, although the precise technique and attendant details will be mentioned in each section, where necessary. Fig. 3. View largeDownload slide Absorption spectra of photoreceptors. (a) rhodopsin, photopsin I, II, III (for human vision); (b) red-light-absorbing form and far-red-light-absorbing form of phytochrome (for photomorphogenesis), (c) chlorophyll a (for photosynthesis), and blue- and UV-light-receptors, cryptochrome, phototropin and UVR8 (for photomorphogenesis). Note that the wavelength with the highest absorption by the human rhodopsin corresponds almost to the lowest absorption by the plant photoreceptors. (Drawn by Satoru Tokutomi). Fig. 3. View largeDownload slide Absorption spectra of photoreceptors. (a) rhodopsin, photopsin I, II, III (for human vision); (b) red-light-absorbing form and far-red-light-absorbing form of phytochrome (for photomorphogenesis), (c) chlorophyll a (for photosynthesis), and blue- and UV-light-receptors, cryptochrome, phototropin and UVR8 (for photomorphogenesis). Note that the wavelength with the highest absorption by the human rhodopsin corresponds almost to the lowest absorption by the plant photoreceptors. (Drawn by Satoru Tokutomi). Light source Photoreceptors mediating fern photomorphogenesis, so far clarified, are phytochromes [30–36], cryptochromes [37], phototropins [38] and neochrome (formerly called phytochrome3) [23]. Both phytochrome and neochrome absorb red light, far-red light, and blue light (Fig. 3b), cryptochromes and phototropins absorb blue light (Fig. 3c). Blue-light-absorbing ZTL/LKP/FKF protein photoreceptors may also be functional in ferns, although their involvement in any phenomenon in ferns has not been reported so far. The chromophores of these photoreceptors are phytochromobilin in phytochrome [39], FADH and pterin in cryptochrome [12] and flavin mononucleotide (FMN) in phototropin and ZTL/LKP/FKF [11]. The chromophores of neochrome have not been studied precisely, but they are undoubtedly phytochromobilin and FMN. Each chromophore absorbs over a wide wavelength range, so that the appropriate light source could be a white fluorescent tube wrapped with red or blue plastic filters. Currently, LED bulbs with a 450- or 480-nm peak for blue light or with a 660-nm peak for red light are also good enough, although LED bulbs are rather expensive for irradiation of large areas. For far-red-light, a LED bulb emitting in around 730 nm is better than an incandescent lamp to reduce the influence of heat from incandescent lamps. Partial cell irradiation with a microbeam Transformation with green fluorescent protein (GFP)-labeled photoreceptor genes enables us to predict the intracellular localization of the photoreceptors. However, multi-functional photoreceptors, for example, phototropins mediating phototropism, chloroplast movement, stomatal opening, leaf flattening, etc. [11], exhibit widely diffused intra-cellular distribution patterns, so that it is hard to determine which GFP localization corresponds to which physiological phenomenon. On the other hand, if a microbeam spotlight illuminating a small part of a cell induces a process, we are sure that the photoreceptive site of this process is present (i.e. in the illuminated area) and the activated photoreceptor is localized there, although we cannot determine the species of photoreceptor involved. Partial cell irradiation can indicate the functional photoreceptor localization associated with each light-induced process. We have made several microbeam irradiators equipped with different functions by arrangement of ordinal light microscopes [40]. An epi-microbeam irradiator using a NIKON Biophot microscope as an optical train of this machine (Fig. 4, [40,41]) is still functional even now after 40 years operating. Fig. 4. View largeDownload slide Schematic diagram of a microbeam irradiator based on an inverted microscope (Biophot, Nikon). LS1 and LS2, the light sources with tungsten-halogen lamps (L; 12 V, 100 W); Ph, photography; Obs, ocular for observation; M, mirror; Pl, polarizer; Dp, depolarizer; HM, half-mirror; D, diaphragm; Id, iris diaphragm; S, shutter; If, interference filter; H, heat-cut filter; W, Wollaston prism; IR, infrared light filter; Fd, field diaphragm; Nd, neutral density filter; C, cut-off filter. The light for microbeam irradiation from LS1 is filtered to provide red, blue, far-red, or infrared light (shown as a red line) through interference and plastic filters, or infrared filters. Then, the light is transformed to a microbeam through a diaphragm unit. The microbeam is reflected by a half-mirror (HM) onto the material for irradiation, but a small part of the microbeam that is not reflected by the HM reaches the ocular for observation or a camera for recording (shown as a red dotted line). The way to go either the ocular or a camera can be selected by changing the light path pulling out a mirror. The material on a stage is observed by infrared light through the infrared-sensitive camera and a monitor screen. To set a microbeam irradiation spot on a material, both the material and an infrared microbeam are shown on the monitor screen and the position is adjusted. Then, the microbeam is changed from infrared to the desired wavelength, be it red, blue, or far-red light, by revolving the filter unit. Yellow lines indicate light beams without filtration except heat-cut filters. Dark red line shows infrared light. Fig. 4. View largeDownload slide Schematic diagram of a microbeam irradiator based on an inverted microscope (Biophot, Nikon). LS1 and LS2, the light sources with tungsten-halogen lamps (L; 12 V, 100 W); Ph, photography; Obs, ocular for observation; M, mirror; Pl, polarizer; Dp, depolarizer; HM, half-mirror; D, diaphragm; Id, iris diaphragm; S, shutter; If, interference filter; H, heat-cut filter; W, Wollaston prism; IR, infrared light filter; Fd, field diaphragm; Nd, neutral density filter; C, cut-off filter. The light for microbeam irradiation from LS1 is filtered to provide red, blue, far-red, or infrared light (shown as a red line) through interference and plastic filters, or infrared filters. Then, the light is transformed to a microbeam through a diaphragm unit. The microbeam is reflected by a half-mirror (HM) onto the material for irradiation, but a small part of the microbeam that is not reflected by the HM reaches the ocular for observation or a camera for recording (shown as a red dotted line). The way to go either the ocular or a camera can be selected by changing the light path pulling out a mirror. The material on a stage is observed by infrared light through the infrared-sensitive camera and a monitor screen. To set a microbeam irradiation spot on a material, both the material and an infrared microbeam are shown on the monitor screen and the position is adjusted. Then, the microbeam is changed from infrared to the desired wavelength, be it red, blue, or far-red light, by revolving the filter unit. Yellow lines indicate light beams without filtration except heat-cut filters. Dark red line shows infrared light. The block of the eyepiece of the Biophot microscope was replaced with a custom-made light source system producing a microbeam of monochromatic light obtained through a series of interference filters coupled with red, blue or far-red plastic filters (Fig. 4, upper part). The system is composed of several units as follows: a halogen bulb as a light source, heat-cut filter, a turret with four holes for red, blue, far-red and infrared filters set in the light path, an electric shutter, two diaphragm units changeable for a selected shape and size of microbeam (Fig. 5), and a complicated light path system with a mirror and two half mirrors to irradiate the sample and simultaneously observe the microbeam on the sample. Image on the ‘Diaphragm unit’, such as the shape of an aperture, is projected on the focal plane. The microbeam shape and position seen on a sample cell on a stage through an eyepiece (Fig. 4, Obs) or on a monitor screen is not a real microbeam light irradiating the sample (i.e. a false figure). It is a composite figure with real sample figure illuminated from the bottom and one of the epi-microbeam lights separated by a half-mirror (Fig. 4, HM) and then reflected three times by mirror (M), PM, and another M and reached the eyepiece (Fig. 4, red dotted line). Therefore, both microbeams on the sample and through the eyepiece should overlap exactly in this system. To test whether the real and false microbeams are positioned in the same area, we can confirm their positions by observation through the eyepiece of the real microbeam spot reflected from a surface-mirror set on the ‘Sample stage’ and the false microbeam spot. An area of 1–2 μm (in practice, 2–3 μm) in diameter can be irradiated by focusing a figure of the diaphragm (hole) on the ‘Sample stage’ where the plant material is set (Fig. 4, ‘Sample stage’). To prepare the appropriate shape and size of the microbeam, two changeable diaphragm units were made (Fig. 5a, b). One unit is made of a rotating disc to select one of six round holes of different sizes (Fig. 5a). The smallest is 1–2 μm in diameter when focused on the sample stage using an objective lens of 100×. If a smaller-sized pinhole is needed, we can make a very small pinhole in a piece of aluminum foil with a very thin needle, and set it on the largest pinhole. The other unit consists of two sets of slits positioned vertically or horizontally relative to each other, with each slit width being changeable by turning a screw (Fig. 5b) so that the shape of the diaphragm can be a square or a rectangle of any size. When we want to irradiate samples with special figures, such as letters or even cartoons, we can draw their figures with black and white patterns on a transparent film and set them at the position of the diaphragm and focus on the sample. The position of a microbeam spot in a microscopic field is not fixed but rather can be moved to any part of the microscopic field by moving the ‘Diaphragm unit’ in a Diaphragm cassette, using fine screws, meaning that any part of a material that is seen in the microscopic field could be irradiated. The excitation light of the microbeam is provided from the top (as epi-illumination) by a light source of a tungsten halogen bulb through interference filters for blue, red, far-red, or infrared light coupled with plastic plates of each color. Each of those is set in four different holes on the rotating disc (Fig. 4, ‘Filter unit’). The irradiation time is controlled by an electrical shutter system (Fig. 4, ‘S’). The plant materials are monitored by infrared light obtained through the infrared filter IR-85 (Hoya, Tokyo, Japan) (Fig. 4, ‘IR’) from the bottom by an infrared-sensitive camera (Fig. 4, ‘Ph’) (C2400–07ER, Hamamatsu Photonics, Hamamatsu, Japan) and observed on a monitor screen. To measure the fluence rate of the microbeam, silicon photodiodes (S1227-66BR, Hamamatsu Photonics, Hamamatsu, Japan) are used. To reduce the light intensity of the microbeam light, we insert neutral-density filters such as ND-3, ND-10, ND-30 and ND-50 (Hoya) in the filter unit [42]. Fig. 5. View largeDownload slide Diaphragm units for pinholes (a) and slits (b). (a) Six pinholes with different sizes are set on a rotating disc. (b) Slit width or length is changeable by means of screws. Fig. 5. View largeDownload slide Diaphragm units for pinholes (a) and slits (b). (a) Six pinholes with different sizes are set on a rotating disc. (b) Slit width or length is changeable by means of screws. Time-lapse recording and observation The start of the photomorphogenesis process could be induced by light irradiation. The time course of the process should be recorded continually and analyzed later for fine understanding of the phenomenon. For this purpose, time-lapse recording under infrared light, that is not absorbed by plants, is indispensable. A system controlling the acquisition of each photographic frame is needed. We use a Leica digital camera (DFC340FX, Leica microsystems, Germany) equipped with Leica Application Suite (Version 2.8.1). Most of the modern digital cameras have their own time-lapse recording system and could be used for this purpose. For precise analyses of photomorphogenesis processes recorded as time-lapse movies we use ‘ImageJ’ software, version 1.33 (shared from http://rsbweb.nih.gov/ij/) [43]. If it is possible, a system for switching the observation light on and off is recommended to avoid heating up the microscope. Cell centrifugation To define the intracellular photoreceptive site for phototropism at the apical part of the protonemata, polarized microbeam experiments are very effective, although they are not sufficient [44]. Because all organelles within the microbeam light path are illuminated simultaneously, it is hard to distinguish which organelles or membrane systems are the real photoreceptive sites. Cell centrifugation has been an excellent technique in cell biology to move organelles down to one side of a cell [45]. When fern protonemata are centrifuged basipetally, most of the organelles move down, but oil droplets move up to the apical part of the cell because they are less dense than water [44,46]. But if oil droplets could be excluded and replaced by the vacuole, the tonoplast would be present besides the plasma membrane. To know which membrane systems are likely to be the photoreceptive sites, two types of centrifuged cells, with or without oil droplets at the apical part, were made by three successive centrifugations, using L-shaped protonema cells induced by phototropism (Fig. 6a, b1⓪) [7,44]. Firstly, the protonema was centrifuged acropetally towards the cell apex to gather the oil droplets at the apical region to the corner of the L-shaped protonema (Fig. 6a①, b1①). Then, the protonema was centrifuged parallel to the basal region of L-shaped protonema acropetally (Fig. 6a②1) or basipetally (Fig. 6a②2) to either direction to chase the oil droplets at the cell corner towards the basal region (Fig. 6b1②) or to gather the oil droplets at the basal region to the corner (Fig. 6b2②). Finally, the protonema was centrifuged parallel to the apical region basipetally (Fig. 6a③) to make a vacant (or vacuolated) apical part (Fig. 6b1③, c1) or an apical part filled with oil droplets (Fig. 6b2③, c2). After partial irradiation with microbeam at the apical part of the protonema, organelles were brought back to the apical part by acropetal centrifugation (Fig. 6a④, b1④, b2④). Fig. 6. View largeDownload slide How to produce a protonema apex without organelles. (a) Directions and order of repeated centrifugations to spin down organelles with or without oil droplets in an L-shaped protonema. (b) The L-shaped protonema was induced by a phototropic response to a red-light source by turning the protonema once (b1⓪). Centrifuge the protonema toward the cell apex to move all oil droplets at the apical part of the protonema to the bent corner (a①, b1①). Centrifuge the protonema parallel to the basal part of L-shaped protonema in either direction acropetally (a②1) or basipetally (a②2) to chase the oil droplets at the corner to the basal direction (b1②) or to move the oil droplets in the basal part to the corner (b2②), respectively. Then, centrifuge the protonema parallel to the apical part of the protonema towards the corner to move the organelles and the nucleus down to make a vacuolated (b1③) or an oil-droplet-filled apex (b2③). After microbeam light irradiation, centrifuge the protonema again to bring the organelles back to the apical part (b1④, b2④). (c) Cross-sections of the sub-apical parts of vacant (1) and oil-droplets-filled protonemata (2) and their magnification views in the right. Bars: 1 μm. Yellow: nuclei, green: chloroplasts, purple: oil droplets. a,c: cited from [24]. Fig. 6. View largeDownload slide How to produce a protonema apex without organelles. (a) Directions and order of repeated centrifugations to spin down organelles with or without oil droplets in an L-shaped protonema. (b) The L-shaped protonema was induced by a phototropic response to a red-light source by turning the protonema once (b1⓪). Centrifuge the protonema toward the cell apex to move all oil droplets at the apical part of the protonema to the bent corner (a①, b1①). Centrifuge the protonema parallel to the basal part of L-shaped protonema in either direction acropetally (a②1) or basipetally (a②2) to chase the oil droplets at the corner to the basal direction (b1②) or to move the oil droplets in the basal part to the corner (b2②), respectively. Then, centrifuge the protonema parallel to the apical part of the protonema towards the corner to move the organelles and the nucleus down to make a vacuolated (b1③) or an oil-droplet-filled apex (b2③). After microbeam light irradiation, centrifuge the protonema again to bring the organelles back to the apical part (b1④, b2④). (c) Cross-sections of the sub-apical parts of vacant (1) and oil-droplets-filled protonemata (2) and their magnification views in the right. Bars: 1 μm. Yellow: nuclei, green: chloroplasts, purple: oil droplets. a,c: cited from [24]. To perform repeated centrifugations to different directions, we designed a special cuvette that consists of two stainless steel rings tightly screwed together. The protonemata were sandwiched between two round glass pieces (18-mm diameter) with a silicon ring spacer in between, and then the cuvette was set into a cassette fitting to a swing-type centrifuge and centrifuged repeatedly towards a desired direction [7,44,47]. This cuvette is also useful to observe gametophytes for long periods without the sample drying out. How to return to the same cell It is not easy to return to the same protonemal cell among many similar cells once it is moved out from the microscopic field, even after only a few minutes; after a few days growth, it is almost impossible. For overcome this issue, we developed a method to monitor a place in the microscopic field using small pieces of glass that are smashed with a mortar and pestle and which are then spread onto the surface of the culture medium. Each piece of broken glass has a unique shape, so that the same piece could easily be found, helping to re-locate the cell of interest near the marker glass piece. Photo-regulation of cell growth Apical growth of protonemal cells The protonemal cells of many fern species, such as A. capillus-veneris and the male fern Dryopteris filix-mas, show apical growth toward the light source under red light for more than a week [48,49], resulting in a long single cell without cell division. Pteris vittata (the Chinese brake fern) is an exception. Protonemata of P. vittata show an apical growth as a long single cell under red light but show neither the polarotropic response perpendicular to the vibration plane of polarized red light nor the phototropism towards a red-light source [7,50], although they do show negative geotropism (personal communication from Michizo Sugai). In A. capillus-veneris, the apical part is filled with organelles, with ~200 chloroplasts within 100 μm of the tip [51]. The center of the nucleus is ~60 μm from the tip of the protonemata in A. capillus-veneris (Fig. 7, [51,52]) compared with 40–50 μm in P. vittata [53]. The protonemal growth rate is reduced when transferred to the dark, this effect occurring more rapidly by a few minutes irradiation with far-red light immediately before the transfer. The far-red light effect was reversed by a red-light pulse; the photoreceptor for this phenomenon was, therefore, predicted to be a phytochrome [31]. The cell growth under red light was inhibited by blue light [35]. Partial cell irradiation with blue light using a microbeam irradiator revealed that the blue light effect for the growth inhibition was found in the nuclear region [35], suggesting that the photoreceptor would be cryptochrome 3 (cry3) or cry4, which is localized in the nucleus (see below) [37]. Fig. 7. View largeDownload slide Light micrographs of longitudinal serial sections of a red-light-grown protonema of A. capillus-veneris. The apical part is filled with organelles. The center of the nucleus is ~60 μm from the tip. The spindle-shaped nucleus has one nucleolus. Reconstruction of the apical part with the serial sections revealed that the cigar-shaped long chloroplasts are attached to the apical dome of the protonema. To section protonemata which are several hundred micrometers long longitudinally, a special arrangement of a protonema-containing block on a microtome sample holder was needed as explained below. The block with a protonema was set so that the protonema was vertical. Then, several sections of the block were cut in the ordinary way with a glass knife, the glass knife was turned through 90 degrees, and the left side of the block on a specimen-holder was cut. When the protonema was viewed from the side, namely from the horizontal direction, with a specifically arranged horizontal microscope, the glass knife was turned back to the ordinal position, some more sections were cut, and the protonema arranged to be parallel to the cut surface; the process was then repeated until the protonema in the block had become exactly parallel to the cut surface of the block, and serial sections were cut. N: nucleus, n: nucleolus. Bar: 10 μm. Cited from [51]. Fig. 7. View largeDownload slide Light micrographs of longitudinal serial sections of a red-light-grown protonema of A. capillus-veneris. The apical part is filled with organelles. The center of the nucleus is ~60 μm from the tip. The spindle-shaped nucleus has one nucleolus. Reconstruction of the apical part with the serial sections revealed that the cigar-shaped long chloroplasts are attached to the apical dome of the protonema. To section protonemata which are several hundred micrometers long longitudinally, a special arrangement of a protonema-containing block on a microtome sample holder was needed as explained below. The block with a protonema was set so that the protonema was vertical. Then, several sections of the block were cut in the ordinary way with a glass knife, the glass knife was turned through 90 degrees, and the left side of the block on a specimen-holder was cut. When the protonema was viewed from the side, namely from the horizontal direction, with a specifically arranged horizontal microscope, the glass knife was turned back to the ordinal position, some more sections were cut, and the protonema arranged to be parallel to the cut surface; the process was then repeated until the protonema in the block had become exactly parallel to the cut surface of the block, and serial sections were cut. N: nucleus, n: nucleolus. Bar: 10 μm. Cited from [51]. At the subapical region, around the base of the dome-shaped apical part of the red-light-grown protonemata, a ring structure, 5–10-μm wide, composed of many microtubules [54,55] and actin microfilaments [56] running perpendicular to the growing axis, was found along the plasma membrane (Fig. 8). The innermost layer of the cell wall at the subapical region consisted of cellulose microfibrils oriented parallel to the microtubules [55]. This microfibril distribution pattern is similar to that found in the innermost layer of seed plant cell walls that control cell shape, cell diameter and the direction of cell growth of seed plant cells [57]. Microfibril distribution might play a significant role in controlling the protonemal cell diameter [55], as in the case of seed plant cells [58]. Hexagonal rosettes of the cellulose synthase complex in the plasma membrane, which can be seen in seed plant cells [58] by freeze-fracture scanning electron microscopy, were also observed at the apical and subapical parts of growing protonemal cells [59], suggesting that the mechanisms of cell wall synthesis in seed plant and fern protonemata might have the same origin. Fig. 8. View largeDownload slide A band of microtubules surrounding the sub-apical part of the protonemata during the polarotropic response induced by turning the vibration plane of the red light at 70°. From left: before the induction of the polarotropic response, 20 min, 1 h and 3 h after induction. Note that the ring structure turned gradually towards the future direction but disappeared once, and then reappeared when the protonema showed bending. Bar: 10 μm. Rearranged from [67]. Fig. 8. View largeDownload slide A band of microtubules surrounding the sub-apical part of the protonemata during the polarotropic response induced by turning the vibration plane of the red light at 70°. From left: before the induction of the polarotropic response, 20 min, 1 h and 3 h after induction. Note that the ring structure turned gradually towards the future direction but disappeared once, and then reappeared when the protonema showed bending. Bar: 10 μm. Rearranged from [67]. Treatment with cytochalasin B (50 μg/ml) destroyed both the actin and microtubule ring structures simultaneously, but colchicine (5 mM) destroyed only the microtubule rings but not the microfilament rings [60]. Furthermore, when apical cell bulging or phototropism of protonemata was induced, the actin ring structure changed its distribution pattern before the microtubule rings did [61](see below). These results suggest that the distribution pattern of microtubules is controlled by the actin ring structure [27]. Overall, the actin rings might control the microtubule rings, which might in turn control the microfibril distribution pattern, although how the location of the actin rings is determined is not known. Phototropism Protonemal cells show positive phototropism towards a light source at the apical part. However, under polarized light, they grow perpendicular to the vibration plane of the light, but not towards the source of polarized light. This phenomenon has been named ‘polarotropism’ [62]. When red-light-grown protonemata were incubated in darkness for 15 h and then irradiated with polarized red light (7 W m−2) vibrating at 45 degrees to the cell axis for 2 min, a polarotropic response could be observed within 30 min of subsequent incubation in the dark [63]. The apical part of the cell started to bulge, probably at the highest absorption of the light, and then clear polarotropism followed [63]. We performed experiments with partial cell irradiation (10-μm wide) for 10 s to find the photoreceptive site for the red polarized light (7 W m−2), and found that the highest effects were found around 5–15 μm from the cell apex, at the base of the apical dome of the protonemata, but not at the very tip of the cell. Next, the center of the cell or cell edge of the photoreceptive site was irradiated and the true photoreceptive site was found to be the cell edge [63], meaning that the vibration plane of the polarized light could be detected when the vibration plane was parallel, but not perpendicular, to the cell wall. To change the growth direction under polarized light, the cell edge of the subapical part of the protonemal cell should be irradiated; however, to continue apical growth under polarized light after the polarotropic response, the photoreceptive site is the very tip of the protonema [64]. Partial cell irradiation with a microbeam is an excellent method, as mentioned above, but has a few issues to which attention should be paid. The microbeam light penetrates deep into the cell and illuminates the entire cell, meaning that the light might illuminate all the organelles and proteins existing inside the beam light path. Hence, to avoid misinterpretation of the results, we performed cell centrifugation to remove all organelles from the photoreceptive site; then, a cell edge of the photoreceptive site with almost no organelles was irradiated with a non-polarized red-light microbeam of 22 W m−2 [44]. After irradiation, the organelles were returned to the cell apex by acropetal cell centrifugation to supply the materials for cell growth. The protonema showed a phototropic response towards the irradiated side [44], revealing that the photoreceptors were localized close to or on the plasma membrane, but not on the centrifuged organelles. The red-light-absorbing receptor was thought to be phytochrome because of the red/far-red light reversibility of this phenomenon [62], but we found that the photoreceptor was not phytochrome but a neochrome [65], formerly called phytochrome 3 (phy3) [66], a chimeric photoreceptor of a phytochrome N-terminus, chromophore-binding domain, and almost the full length of phototropin [17]. The ring structure, composed of microtubules and microfilaments around the subapical part of the protonemata, showed a drastic change during the phototropic response (Fig. 8). Before a tropistic response or a slight change in cell shape could be detectable at the apical dome region, the actin rings and the microtubule rings had already changed their distribution. Where the final resulting curvature was small, the ring structure changed its distribution gradually towards the bending direction, but when the resulting curvature was very sharp, that is, the protonema had to bend very sharply, the ring structure tentatively disappeared as a result of the actin filaments and microtubules depolymerizing, and then reappeared again after the cell bending had been accomplished [67]. When only one side of the protonemal apical dome was irradiated with the red-light microbeam, the ring structure at the irradiated side disappeared and reappeared after the curvature had been accomplished (Fig. 9) [67]. Fig. 9. View largeDownload slide Schematic illustration showing the dynamic changes of a ring structure during phototropic responses. When sharp bending was induced, the ring structure disappeared once and reappeared after the protonemal bending. When one side of the protonemata was irradiated with red light to induce a phototropic response, the ring at the irradiated region disappeared transiently and reappeared after bending. Fig. 9. View largeDownload slide Schematic illustration showing the dynamic changes of a ring structure during phototropic responses. When sharp bending was induced, the ring structure disappeared once and reappeared after the protonemal bending. When one side of the protonemata was irradiated with red light to induce a phototropic response, the ring at the irradiated region disappeared transiently and reappeared after bending. Apical bulging When red-light-grown protonemata were irradiated with white or blue light (1.2 W m−2) continuously, the apical dome of the protonemata started to bulge within 2 h of irradiation [55,68]. Partial cell irradiation at the apical dome was effective for the induction of bulging [36], meaning that the apical dome is the photoreceptive site. Polarized-blue light vibrating parallel to the cell axis was more effective than light vibrating perpendicularly for achieving apical bulging, suggesting that the photoreceptor was localized close to the plasma membrane at the apical dome of the protonemata, with the transition moment parallel to the membrane [36]. Thirty minutes after irradiation to induce the apical bulging, the actin ring disappeared, followed by the microtubule ring [61]. Then, the direction in which the innermost microfibrils ran became random, and the cell bulging could be recognized [55]. Apical bulging is the first observable phenomenon of the transition from 1D to 2D growth of fern gametophytes, an effect which spreads perpendicular to the direction of the incident light, resulting in flat patterns for easy and effective light capture [69]. Hence, to know how the cell becomes flat, and whether there is any specific part played by cell extension, protonemata growing upward on an agar surface were irradiated with white light from the top to induce apical bulging and subsequent cell division (Fig. 10a, b). Then, small active carbon grains were placed at appropriate intervals along the equatorial position of the bulbous apical cells (Fig. 10a, c). After continuous white light irradiation from a horizontal direction to induce apical flattening, the distances between the carbon grains were measured (Fig. 10d). Contrary to the prediction, that the irradiated region might expand more than the shaded side, the cell wall expanded uniformly around the equatorial area of the bulbous cells (Fig. 10e) [69]. The mechanism by which the apical cell becomes flattened remains a target for future studies. Fig. 10. View largeDownload slide How to induce flattening of the apical cell of the protonema and how to measure the growing region of the apical cell. (a) Schematic illustration of experimental procedures. (b) The time schedule of the light irradiation. A protonema grew on the agar surface towards the red-light source (R1), then grew up towards R2. White light (W1) was shone from above to induce cell division. After cell division, small grains of active carbon were put on the equatorial part of the apical cell. Then, white light (W2) was shone from the horizontal direction to induce cell flattening. (c) Top views of the apical cell with carbon grains before and after cell flattening. Bar: 20 μm. (d) The distances between the adjacent grains were measured before W2 irradiation and after cell flattening and the relative rates were calculated with the equation shown below. (e) The relative rates of cell growth during W2 irradiation were plotted. Note that the rate of cell wall growth around the equatorial area of the apical dome was approximately constant. a,b,c,d redrawn from [69], e, cited from [69]. Fig. 10. View largeDownload slide How to induce flattening of the apical cell of the protonema and how to measure the growing region of the apical cell. (a) Schematic illustration of experimental procedures. (b) The time schedule of the light irradiation. A protonema grew on the agar surface towards the red-light source (R1), then grew up towards R2. White light (W1) was shone from above to induce cell division. After cell division, small grains of active carbon were put on the equatorial part of the apical cell. Then, white light (W2) was shone from the horizontal direction to induce cell flattening. (c) Top views of the apical cell with carbon grains before and after cell flattening. Bar: 20 μm. (d) The distances between the adjacent grains were measured before W2 irradiation and after cell flattening and the relative rates were calculated with the equation shown below. (e) The relative rates of cell growth during W2 irradiation were plotted. Note that the rate of cell wall growth around the equatorial area of the apical dome was approximately constant. a,b,c,d redrawn from [69], e, cited from [69]. Photocontrol of cell division Photoreceptors and their localization Red-light-grown single-celled protonemata divide when they are transferred to white light conditions [48]. Partial cell irradiation with a microbeam of blue light (30-μm wide, 3.6 or 6.7 W m−2, 2 min) revealed that the photoreceptive site for the induction of cell division was ~60 μm from the protonemal tip, which corresponded to the nuclear region [70]. If a protonema was centrifuged along the cell axis to spin its nucleus down in the basipetal direction, the photo-sensitive site for cell division also moved down to the new nucleus region [46]. Considering that the photoreceptive site for blue light for the apical cell bulging effect was localized on the plasma membrane at the apical dome [36], the blue-light receptors for the two phenomena must be different, even though the blue-light responses occurred simultaneously in the same cell. This was the first report at that time indicating the separation of different intracellular photoreceptive sites between two independent phenomena in one cell. The photoreceptor controlling cell division is thought to be one of the cryptochromes, either cry3 or cry4, or both because of their translocation from the cytosol to the nucleus [37]. Since phototropins are localized on the plasma membrane in A. thaliana [71], the blue-light receptor that mediates apical bulging of fern protonemata might be either phototropin 1 (phot1) and/or phot2. Molecular biological techniques are not yet developed in ferns and, thus, investigation of a phot1 phot2 double knockout mutant line of A. capillus-veneris is not yet possible, although we have screened the phot2 mutant line in this fern species [38]. The phot2 mutant still showed apical bulging, suggesting that phot1 is at least one of the phototropins involved in the apical bulging (data not published). Another interesting but unresolved issue in fern gametophytes is why red- and blue-light effects on cell division are opposite in spore germination and in protonemata. In the former case, red light promotes cell division while blue light inhibits it, whereas blue light induces cell division while red light inhibits it in the latter [72]. Cell cycle Not only cell division but also each step of the cell cycle in fern protonemata is controlled by blue light as well as red light. Under continuous red light, the cell cycle stays at the beginning of the Gap1 (G1) phase, but blue light irradiation or transfer into darkness restarts the progress of the cell cycle [73]. The duration of the G1 phase is controlled by the blue-light intensity, G1 being shortened by higher intensity and lengthened by lower intensity light [29,74]. The photoreceptor(s) involved have not yet been identified, but are predicted to be either cry3 or cry4 [37]. The G1 stage progressing in the dark could be reverted to the early G1 phase by red light irradiation. This red light-effect could be canceled out by far-red light irradiation, indicating the involvement of phytochrome [75] and meaning that phytochrome maintains the cell cycle of fern protonemata at the beginning of the G1 phase under continuous red light. The G2 phase is also under the control of phytochrome, the duration of G2 lengthening under far-red light, although this effect can be reversed by red light [29,74]. M phase (The mitotic (M) phase), however, was not light controlled [29]. It is questionable why the fern gametophyte cell cycle should be controlled so precisely by light, whether these physiological controls are crucial, and what kind of genes have their expression mediated by red or blue light. All of these questions need to be answered. Position of cell division Cell division occurs at some distance from the tip of long single-celled protonemata, ~30–50 μm under blue light and ~80–110 μm in darkness in A. capillus-veneris [49,52,76]. The position of cell division is associated with the location of the pre-prophase band (PPB) of microtubules on the plasma membrane [49,76], as occurs in angiosperm cells [77,78]. The microtubule structure appears only during pre-prophase at the future site of the cell plate attachment, but usually disappears at the boundary of prophase/prometaphase before cell plate formation. It is not known how the position of PPB is regulated, although some components found in cell division zone are remained through out the cell division stages, as possible memories of the former PPB site [79]. Since the PPB appears near the nucleus in general, the association between the location of the nucleus and the PPB formation site was tested in long protonemal cells [80]. Red-light-grown straight protonemal cells or L-shaped cells were centrifuged towards the protonemal base to bring down the nucleus at various time intervals after the induction of cell division by blue light (Fig. 11a). Then, the PPB was visualized by immunofluorescent staining using anti-tubulin antibody and a fluorescein-conjugated secondary antibody. PPB was observed only at the new nuclear position when centrifugation took place before PPB formation (Fig. 11a2), and only at the former position when centrifugation was performed after PPB formation (Fig. 11a4), and at both nuclear positions when centrifugation took place during PPB formation (Fig. 11a3) [80]. Fig. 11. View largeDownload slide Schematic illustrations of the PPB formation, destruction, and reassembly by cold treatment in centrifuged cells. (a) The stage of nuclear cycle that is effective on PPB formation. (1) PPB formation without centrifugation. PPB was formed near the nuclear region. (2) Centrifuged before prophase. PPB was formed at the new nuclear region. (3) Centrifuged during PPB formation at prometaphase to metaphase. Two PPBs could be formed at the original nuclear region as well as at the new nuclear region. (4) Centrifuged after metaphase. PPB was not formed at the new nuclear region. (b) Reassembly of the PPB at the original nuclear position. A protonema was centrifuged after PPB formation (1, 2) at 25°C, and then chilled at 0°C for 5 min to depolymerize the PPB (3). Warming up the protonemata at 25°C induced reassembly of the PPB at the former PPB site, although the centrifuged nucleus was still far down from the PPB (4). (c) The sites of PPB formation following cell division in the protonemata with colchicine-induced two nuclei. 1, 2, non-centrifuged cells with two nuclei parallel (1) or perpendicular (2) to the orientation of the PPB. 3, 4, centrifuged cells with two associated (3) or separate nuclei (4). If the two nuclei were separated by centrifugation, two PPBs were formed (4). (d) The stage of nuclear cycle that is effective on PPB depolymerization. Two PPBs were induced at different parts of the protonema by nuclear relocation as a result of centrifugation during the timing of PPB formation, then the nucleus was sent back to its original position at (1) prophase, (2) prophase and prometaphase transition, and (3) prometaphase to telophase transition. PPB depolymerization was induced by prometaphase or telophase nuclei. a,b,c,d are redrawn from [80–82,84], respectively. Fig. 11. View largeDownload slide Schematic illustrations of the PPB formation, destruction, and reassembly by cold treatment in centrifuged cells. (a) The stage of nuclear cycle that is effective on PPB formation. (1) PPB formation without centrifugation. PPB was formed near the nuclear region. (2) Centrifuged before prophase. PPB was formed at the new nuclear region. (3) Centrifuged during PPB formation at prometaphase to metaphase. Two PPBs could be formed at the original nuclear region as well as at the new nuclear region. (4) Centrifuged after metaphase. PPB was not formed at the new nuclear region. (b) Reassembly of the PPB at the original nuclear position. A protonema was centrifuged after PPB formation (1, 2) at 25°C, and then chilled at 0°C for 5 min to depolymerize the PPB (3). Warming up the protonemata at 25°C induced reassembly of the PPB at the former PPB site, although the centrifuged nucleus was still far down from the PPB (4). (c) The sites of PPB formation following cell division in the protonemata with colchicine-induced two nuclei. 1, 2, non-centrifuged cells with two nuclei parallel (1) or perpendicular (2) to the orientation of the PPB. 3, 4, centrifuged cells with two associated (3) or separate nuclei (4). If the two nuclei were separated by centrifugation, two PPBs were formed (4). (d) The stage of nuclear cycle that is effective on PPB depolymerization. Two PPBs were induced at different parts of the protonema by nuclear relocation as a result of centrifugation during the timing of PPB formation, then the nucleus was sent back to its original position at (1) prophase, (2) prophase and prometaphase transition, and (3) prometaphase to telophase transition. PPB depolymerization was induced by prometaphase or telophase nuclei. a,b,c,d are redrawn from [80–82,84], respectively. Reassembly of PPB after cold treatment is also interesting (Fig. 11b). When a protonema with PPB at the apical part (Fig. 11b1) was centrifuged to bring down the nucleus (Fig. 11b2), then chilled at 0°C to depolymerize the PPB (Fig. 11b3) and subsequently warmed up to 25°C, a PPB was assembled at the original PPB position but not at the new nuclear region (Fig. 11b4), indicating that the memory of the PPB was maintained during the cold treatment [81]. Treatment with 4-mM caffeine inhibits cell plate formation, resulting in a cell with two nuclei [82]. When the protonema with two nuclei was centrifuged and if the two nuclei stayed separate, two PPBs appeared around each nucleus (Fig. 11c) [83]. These results confirm that PPB is formed at the nuclear region. Nuclei are also involved in PPB disassembly [84]. When protonemata were centrifuged at 12 h after the induction of cell division by blue light to move the nucleus downwards, PPB appeared either at the former and/or the new nuclear position in the protonemata, as mentioned above (Fig. 11d). Then the nucleus was returned to its original position by centrifugation. If the nucleus at prometaphase (Fig. 11d2) or telophase (Fig. 11d3) was taken back to the PPB at the original position, the PPB disappeared (Fig. 11d2, 3); however, the prophase nucleus did not show such an effect (Fig. 11d1). The PPB formed at the new nuclear position was not disrupted if its nucleus was taken back to the original position (Fig. 11d2). These results suggest that for PPB formation and PPB disruption, a nucleus at a specific stage and/or some factor(s) in the endoplasm that co-centrifuged with the nucleus must be needed. Direction of cell division Plant cells that are surrounded with rigid cell walls attach to each other tightly, so that the position of each cell cannot be changed once established. Hence, the site and direction of cell division should be controlled precisely, predicting the future shape of the tissue. Fern prothallia spread perpendicularly to the incident light [85–88]. In A. capillus-veneris protonemata, the first step to two-dimensional growth for prothallia occurs at the third cell division, that is, the first longitudinal division at which cell plate becomes parallel to the direction of the incident light [88]. Under polarized white light, the fern gametophytes spread parallel to the vibration plane of the polarized light irrespective of the direction of the light [85], and the cell plate of the first cell division for transition to two-dimensional growth also occurs mostly at the third division, perpendicular to the vibration plane [85]. The regulatory mechanisms of the direction change of cell plate from perpendicular to parallel to the protonemal axis, that is, the transition point from one-dimensional to 2D growth are not known. It might depend on either the direction of incident light, the vibration plane of the polarized light, or on other factors. In any case, it is clear that the cell shape becomes flattened, perpendicular to the future cell plate of the first longitudinal division (Fig. 12) [85,88], indicating that the cell shape may decide the direction of cell division as shown recently in A. thaliana [89], although the precise mechanism is not known. Fig. 12. View largeDownload slide Direction of the cell division and prothallial flattening. An early stage of prothallial growth was observed from vertical and horizontal directions. (a) To take photographs of a protonema repeatedly from the top and the side, a special cuvette (2 × 2 × 5 mm) was made from a transparent thin plastic plate. A sterilized spore was embedded in the agar medium, and a protonema was cultivated on the surface of the agar medium with horizontally irradiated red light. Then, the protonema was cultivated under white light, being continuously irradiated from the top. Photographs were taken frequently as a top view or a side view by turning the cuvette through 90°. (b) Serial photographs of top and side views of the protonema cultivated in the plastic cuvette. The apical cell became flattened and the first longitudinal division occurred parallel to the incident white light at the third cell division. Bar: 50 μm. b, cited from [69]. Fig. 12. View largeDownload slide Direction of the cell division and prothallial flattening. An early stage of prothallial growth was observed from vertical and horizontal directions. (a) To take photographs of a protonema repeatedly from the top and the side, a special cuvette (2 × 2 × 5 mm) was made from a transparent thin plastic plate. A sterilized spore was embedded in the agar medium, and a protonema was cultivated on the surface of the agar medium with horizontally irradiated red light. Then, the protonema was cultivated under white light, being continuously irradiated from the top. Photographs were taken frequently as a top view or a side view by turning the cuvette through 90°. (b) Serial photographs of top and side views of the protonema cultivated in the plastic cuvette. The apical cell became flattened and the first longitudinal division occurred parallel to the incident white light at the third cell division. Bar: 50 μm. b, cited from [69]. Branching Red-light-grown protonemata were irradiated with white light to induce cell division, then centrifuged basipetally to relocate the nucleus to a location ~200–250 μm from the apex, and then kept under polarized red light. After cell division, a nucleus in the apical cell returned to the original cell apex position and restarted cell growth if the distance of nuclear relocation was rather short. However, when the cells divided rather far from the original apex, branch formation near the septum occurred frequently [90] (Fig. 13). Furthermore, if the divided protonemata were centrifuged basipetally to relocate the new upper nucleus down to the septum, the frequency of branch formation increased. On the other hand, if cells were centrifuged acropetally, the percentage of cells growing at the original apex increased. These results indicate that either side of the cell, the original apex or the newly established branch apex, can be an growing apex. Considering that the large and spindle-shaped nuclei in these narrow cells might not be able to change their direction (or to rotate) during the cell growth and centrifugation, either side of the nuclei could become a front or rear face in apically growing cell, indicating that the nuclei might not have physiological polarity. When we centrifuged the branched cells basipetally or acropetally, either ends of the old or new apices could become a new apex of the cells [90]. These phenomena suggest that the polarity of the protonemal cells could easily be reversed by nuclear positioning as a consequence of cell centrifugation. The nuclear behavior during branch formation is interesting in relation to cytoskeletal patterns, refer to reviews for more details [90,91]. Fig. 13. View largeDownload slide Branching of protonema induced by cell centrifugation. Red-light-grown protonemata were irradiated with white light to induce cell division, and then centrifuged basipetally following polarized red light irradiation as shown in the time schedules with or without incubation in the dark (a). (a) A photograph of a protonema with two branches near the septum. (b) The results of this experiment showed that branches were induced frequently near the septum. (c) The branched protonema could grow at the new or the old apex if the nucleus was brought to either apex by centrifugation. Two nuclei were shown as a result of 4′,6-diamidino-2-phenylindole (DAPI) staining. The way to induce the apical growth on either apex is shown in the illustration on the right. Cited from [90]. Fig. 13. View largeDownload slide Branching of protonema induced by cell centrifugation. Red-light-grown protonemata were irradiated with white light to induce cell division, and then centrifuged basipetally following polarized red light irradiation as shown in the time schedules with or without incubation in the dark (a). (a) A photograph of a protonema with two branches near the septum. (b) The results of this experiment showed that branches were induced frequently near the septum. (c) The branched protonema could grow at the new or the old apex if the nucleus was brought to either apex by centrifugation. Two nuclei were shown as a result of 4′,6-diamidino-2-phenylindole (DAPI) staining. The way to induce the apical growth on either apex is shown in the illustration on the right. Cited from [90]. Chloroplast photorelocation movement For precise analytical studies on the mechanisms of photo-responses, simple and spatiotemporally short and rapid processes would be ideal. Chloroplast photorelocation movement is one of the best candidate processes in that sense because de novo mRNA synthesis is not involved in the signal transduction pathways [92], the phenomenon is cell autonomous [93], and the response occurs within a few minutes of light irradiation [47]. Fern gametophytes are an excellent material for such studies because they are not surrounded by tissue and, therefore, microbeam irradiation and observation of the response are very easy. Chloroplasts gather at the place irradiated with weak light to increase the efficiency of photosynthesis (accumulation response or weak light response, [94]), and move away from strong light to avoid chloroplast photo-damage (avoidance response, or strong light response, [95]). Chloroplast movement is thought to be important for plants living under canopies where fluctuation of the light intensity happens frequently, but the ecological importance of this phenomenon for plants living under direct sunlight for the entire day is not well understood. It has recently been found that under such conditions, most chloroplasts stay in the chloroplast-avoidance position on the anticlinal walls of long thin palisade cells, even under low light conditions, meaning that chloroplast relocation movement is not functional in these leaves [96]. How to observe chloroplast movement The speed of the chloroplast movement is very low, <1 μm/min [97–100], so that it is hard to detect the movement by the naked eye, even with the aid of a microscope. In former times, the movement was mainly detected only by comparing the distribution patterns before and after light irradiation [101]. Thanks to technological advancements, new methods to detect the moving chloroplasts are now available. The time course of red-light transmittance or absorption changes detected spectrophotometrically through a leaf is a well-established conventional method [40,102–105]. However, in small fern gametophytes, the method is not available, but detecting a real behavior (or movement) of each chloroplast is easy by taking photographs every minute by time-lapse photography and analyzing them on a monitor screen [40]. For this experiment, partial cell or partial chloroplast irradiation with the microbeam spot is quite effective [41,99,106]. Various optical instruments, including laser scanning microscopes, are now available for us to record the behavior of each targeted chloroplast continuously and to analyze it precisely [40]. Chloroplast behavior The shape and size of chloroplasts are dependent on the plant species. In mosses, ferns, and angiosperms, many disc-shaped (or lens-shaped) chloroplasts are found in one cell and their size is roughly 5 μm in diameter [99,107,108]. Smaller-sized chloroplasts move more rapidly than large ones [109]. On the other hand, Mougeotia (Charophyta), Selaginella (Lycopodiophyta) and Klebsormidium (Charophyta) are exceptions as they have a single large chloroplast in each cell [101], although they still show chloroplast movement [110]. Microbeam experiments using A. capillus-veneris gametophytes revealed that chloroplasts slide but do not roll in any direction without rotating in both accumulation and avoidance responses because chloroplasts attach their concave side to the plasma membrane, meaning that they do not have polarity for movement [40,99,111]. The velocity of chloroplast movement differs between the accumulation and avoidance responses. When a part of the cell of the A. capillus-veneris gametophytes was irradiated either continuously or with a short pulse from a microbeam of blue or red light to induce accumulation responses, chloroplasts moved towards the irradiated spot at a constant speed even though the intensity of the microbeam light was different [97,99]. In contrast, the velocity of chloroplasts upon avoidance response induced by microbeam irradiation was dependent on light intensity; the higher the light intensity, the faster the chloroplast escaped from the light beam [96–99]. In any case, the velocity of the chloroplast movement is low, around 1 μm per minute on average in both cases [97,99]. Photoreceptive sites and photoreceptors Microbeam irradiation with red or blue light of a small part (e.g. 10-μm diameter) of a dark-adapted and largely vacuolated A. capillus-veneris gametophyte cell, in which most of chloroplasts were at the anticlinal walls so that no chloroplast was at the irradiated site, induced the accumulation response, meaning that the photoreceptors were localized on or close to the plasma membrane [106]. In contrast, chloroplasts showed the avoidance response only when illuminated with strong blue light [42], or, more effectively, when a chloroplast was partially irradiated [99], suggesting that the photoreceptors were localized on the chloroplasts. The effective wavelength to induce chloroplast movement is blue light in general, but red light is also effective in many algae, mosses and ferns for accumulation responses [101,112–114]. In recent years, many mutant lines defective with respect to chloroplast movement have been isolated mainly in Arabidopsis (for reviews, [114–116]) but also in A. capillus-veneris [17,38]. Analyses of these lines revealed that the blue-light-receptor phot1 was responsible for the accumulation response [117], while phot2 was responsible for the avoidance as well as accumulation responses [38,118,119]. In Arabidopsis leaves, phot2-GFP was detected on the envelope of chloroplasts as well as on the plasma membrane, while phot2 was also detected biochemically on the isolated chloroplast membrane [120]. Phototropins have two light, oxygen, voltage (LOV) domains at the N-terminal half. The LOV domains consist of ~100 amino acid residues to which FMN is covalently bound. The C-terminal half is a serine/threonine kinase [121]. Besides chloroplast movement, phototropins mediate phototropism [122], stomatal opening [123], leaf flattening [71,117], leaf development [124], and growth inhibition of hypocotyls [125]. These phenomena play important roles in photosynthesis efficiency at least in A. thaliana. As a red-light-receptor, A. capillus-veneris has a neochrome, a chimeric photoreceptor of phytochrome and phototropin as mentioned above [17,66]. Another neochrome with a different origin but with a structure similar to that of the Adiantum neochrome was found in the filamentous green alga Mougeotia [65,126]. Surprisingly, the Mougeotia neochrome was functional in A. capillus-veneris gametophytes [65], suggesting that the red-light receptor that Wolfgang Haupt had predicted to be a phytochrome in the regulation of Mougeotia chloroplast movement [127] might be this neochrome. A phylogenetic study of phytochromes, including neochrome, was performed using a wide range of plant groups and revealed that the Adiantum neochrome originated in the bryophyte hornwort and was acquired by ferns by horizontal gene transfer [126,128]. Signal transduction pathways The signal transferred from photoreceptors to chloroplasts is not known. Chloroplast movement is cell autonomous and starts within a few minutes after light irradiation, so that the signal transduction pathway would be simple. However, no reliable data have been shown despite a long history of studies (see review [129]). In this situation, several experiments to identify the characteristics of the signal or signal transduction pathway were performed by taking advantage of fern gametophyte experimental technology, which includes various techniques such as microbeam irradiation and cell ligation [26,130]. Red-light-grown, long, single-celled protonema cells divide at the apical part of the cell in response to white light, resulting in a short apical cell and a long basal cell. Once the cell division occurs, the long basal cell becomes more sensitive with respect to chloroplast movement than does the apically growing cell without cell division [131]. To determine whether de novo mRNA synthesis is necessary in the signal transduction pathway or not, a basal cell-fragment without a nucleus was prepared [92]. The part of the cell below the nuclear region was ligated with a thin thread, and the upper nuclear region of the cell was cut-off with a hand-made pair of scissors. In the remaining enucleated cell fragments, both chloroplast accumulation and avoidance responses could be induced by partial cell irradiation with red- or blue-light microbeams or by whole-cell irradiation with polarized red or blue light, indicating that de novo mRNA synthesis was not necessary for the signal transduction involved in chloroplast movement [92]. How do chloroplasts decide in which direction to move? Chloroplasts must detect either the direction of signal flows or simply a higher concentration of signals. When two red-light microbeams with the same fluence rate were used simultaneously to illuminate the central part of a dark-adapted A. capillus-veneris gametophyte cell (Fig. 14), the chloroplasts around the cell periphery at the same distance from these microbeams (see Fig. 14 a2, b) moved to the center between the two microbeams but not to either beam spot. This result indicates that the chloroplasts detected the higher concentration of signals but not the direction of the signal flow [132]. Fig. 14. View largeDownload slide Chloroplast movement induced by simultaneous irradiation with two microbeams in A. capillus-veneris gametophytes. Two red-light microbeams (4 μm in diameter, 20 μm apart) with the same fluence rates (a, 10 W m−2) or different fluence rates (c, 25 and 10 W m−2) were switched on simultaneously on different parts of dark-adapted gametophyte cells. Pathways of chloroplast movement are shown in the photographs a3 and c3 and in the schematic illustrations b and d. Photographs shown before (a1, c1), during (a2, c2) irradiation, and after (a3, c3) movements. Note that chloroplasts 1 and 2 moved between the two microbeams when the beams had the same fluence rate (a3, b), but both chloroplasts 1 and 2 moved towards the spot with the higher fluence rate when the beams had different fluence rates (c3, d), suggesting that chloroplasts move to the higher concentration of signals but not to the spot raising the signals. Bars: 10 μm. Cited from [132]. Fig. 14. View largeDownload slide Chloroplast movement induced by simultaneous irradiation with two microbeams in A. capillus-veneris gametophytes. Two red-light microbeams (4 μm in diameter, 20 μm apart) with the same fluence rates (a, 10 W m−2) or different fluence rates (c, 25 and 10 W m−2) were switched on simultaneously on different parts of dark-adapted gametophyte cells. Pathways of chloroplast movement are shown in the photographs a3 and c3 and in the schematic illustrations b and d. Photographs shown before (a1, c1), during (a2, c2) irradiation, and after (a3, c3) movements. Note that chloroplasts 1 and 2 moved between the two microbeams when the beams had the same fluence rate (a3, b), but both chloroplasts 1 and 2 moved towards the spot with the higher fluence rate when the beams had different fluence rates (c3, d), suggesting that chloroplasts move to the higher concentration of signals but not to the spot raising the signals. Bars: 10 μm. Cited from [132]. If that is the case, chloroplasts may have signal sensors around the entire peripheral region. To determine how accurately chloroplasts detect the signal, and whether they detect the difference between or the ratio of fluence rates, two adjacent parts (each 50 μm in width) of a long protonemal cell were irradiated with various combinations of two red-light microbeams with high and low ratios (H/L) (1:1.2, 1:1.5, 1:2 and 1:10) with different light intensities (1, 0.1 and 0.01 W m−2; Fig. 15, [7,25]). The chloroplasts moved to the site irradiated with the higher-intensity light if the H/L ratio was >2, even though the light intensity was very low, such as 0.007 W m−2, but movement did not depend on the difference in fluence rate. Fig. 15. View largeDownload slide Chloroplasts sense signals as high/low ratio of fluence rates rather than the difference between them. (a) Various sets of different fluence rates (high/low ratios or differences) of red light were made and were shone on the adjacent parts of a long protonemal cell. (b) Then, to which part of the cell the chloroplasts moved is summarized in the table. Chloroplasts sense the ratio of fluence ratios rather than the difference. Redrawn from [25]. Fig. 15. View largeDownload slide Chloroplasts sense signals as high/low ratio of fluence rates rather than the difference between them. (a) Various sets of different fluence rates (high/low ratios or differences) of red light were made and were shone on the adjacent parts of a long protonemal cell. (b) Then, to which part of the cell the chloroplasts moved is summarized in the table. Chloroplasts sense the ratio of fluence ratios rather than the difference. Redrawn from [25]. Characteristics of the signal The speed of intracellular signal transfer in the light-induced chloroplast accumulation response was calculated using dark-adapted protonemal cells [132]. A part of the long basal cells of the protonemata was irradiated with a 1-W m−2 red-light microbeam 20 μm in width for 1 min, then the time at which each chloroplast at some distance from the microbeam started moving after the microbeam was switched off was measured based on time-lapse photographs taken every minute [133]. The calculated data indicated that the speed was dependent on the cell polarity of the protonemata. The mean speed of the signal transmission was 2.3 μm min−1 from the base to the tip and 0.6 μm min−1 from the tip to the base. In prothallial cells, where polarity was not clear, the speed of the signal transmission was ~1–1.5 μm min−1 on average in any direction [133,134]. More interestingly, the signal could be transmitted at the same speed through a narrow space between a chloroplast and the plasma membrane, to which the chloroplast attached firmly as an obstacle blocking the pathway of the signal from the microbeam-irradiated area to the targeted chloroplast (Fig. 16) [100]. We further studied the temperature-dependency of the signal transmission using a custom-made microbeam-irradiator with a custom-made temperature-controlled stage. The speed of the signal transmission proved to be temperature-dependent within the 15°C–30°C range tested, the speed approximately doubling in response to a 10°C increase, suggesting close agreement to the Arrhenius equation in chemical reactions. This indicates the involvement of chemical reactions rather than the simple diffusion of chemicals such as calcium ions [100]. Fig. 16. View largeDownload slide Signals for accumulation response could be transferred through the narrow space between the chloroplasts and the plasma membrane. A part beside a chloroplast that attached the plasma membrane as an obstacle was illuminated with a red microbeam to induce an accumulation response in other chloroplasts staying at the same distance from the microbeam on the same side of the microbeam and beyond the obstacle chloroplast in a dark-adapted gametophyte cell. The paths through which the chloroplasts moved toward the microbeam were traced (a, white lines) and their speeds of signal transfer were calculated (b). Note that both chloroplasts moved straight to the microbeam without making a detour and the speeds of signal transfer were the same in both chloroplasts, indicating that the signal passed under the obstacle chloroplast without delay. Bar: 10 μm. Cited from [100]. Fig. 16. View largeDownload slide Signals for accumulation response could be transferred through the narrow space between the chloroplasts and the plasma membrane. A part beside a chloroplast that attached the plasma membrane as an obstacle was illuminated with a red microbeam to induce an accumulation response in other chloroplasts staying at the same distance from the microbeam on the same side of the microbeam and beyond the obstacle chloroplast in a dark-adapted gametophyte cell. The paths through which the chloroplasts moved toward the microbeam were traced (a, white lines) and their speeds of signal transfer were calculated (b). Note that both chloroplasts moved straight to the microbeam without making a detour and the speeds of signal transfer were the same in both chloroplasts, indicating that the signal passed under the obstacle chloroplast without delay. Bar: 10 μm. Cited from [100]. Careful observation of microbeam-induced chloroplast movement showed that chloroplasts were always monitoring the signals from photoreceptors, at least in the accumulation response. In other words, signals were released continuously from activated photoreceptors. Soon after the first pulse of the blue microbeam, the chloroplast started moving toward the beam-irradiated region, but some time after, if a second microbeam was used to illuminate a different part of the cell, the chloroplast changed direction from the first microbeam to the second [111]. Neochrome could be activated or inactivated by red or far-red light, respectively. When chloroplast movement as part of the accumulation response and mediated by neochrome was induced by a red microbeam, and then the entire cell was irradiated with far-red light to inactivate the neochrome, the chloroplast stopped moving after some lag time (presumably, the period necessary for the signal released from the photoreceptor to reach the chloroplast), but restarted when the second red microbeam was switched on, indicating that the chloroplast stopped when the signal from neochrome ceased, but restarted when the new signals from the neochrome reached the chloroplast [132]. The signals from neochrome and phototropin must be the same, although this has not yet been proven. Lifetime of the signals The signals released from photoreceptors for the accumulation and avoidance responses are different. When part of an A. capillus-veneris prothallial cell was irradiated with a microbeam (of either blue or white light) strong enough to induce an avoidance response, the chloroplasts found within the beam moved away, but the chloroplasts outside the beam moved towards it [42], indicating that the signal for the avoidance response is transmitted over only a short distance, whereas that for the accumulation response can be transmitted over a relatively long distance. The lifetime (or duration of the active form) of the signal for the accumulation response is different from that for the avoidance response. When the strong microbeam was switched off, the chloroplasts outside the beam rushed into the formerly beam-irradiated area as a result of the accumulation response [42,134], indicating that, as soon as the microbeam was switched off, the signal for the avoidance response fell quickly below the threshold value, whereas the signal for the accumulation response was still active and sufficient for the induction of chloroplast movement, so that the accumulation response overcame the avoidance response. The lifetimes were calculated twice by different authors in A. capillus-veneris gametophyte cells and were 3–4 and 6 min for the avoidance response and 19–28 and 30–40 min for the accumulation response [42,106,134]. Mechanism of light-mediated chloroplast movement Fern gametophytes are extremely useful experimental systems for use in cell biology, especially a photobiological point of view, as described above. For genetic analyses, however, they are not good materials; mutants are readily obtained due to their haploid nature, as long as the target genes are not duplicated, but subsequent gene analysis is almost impossible because of the large genome size. On the other hand, we have selected mutants with altered chloroplast movement in Arabidopsis and have clarified the roles of many genes involved in this process. Please refer to review articles about the precise mechanisms by which chloroplasts move, as revealed in A. thaliana [47,116,135]. Here, I will summarize the results obtained in A. capillus-veneris. We found that in Arabidopsis, chloroplast movement involved a novel actin structure located between the chloroplast and the plasma membrane named chloroplast actin filaments (abbreviated to ‘cp-actin filaments’) [136,137]. Under the steady state, during which chloroplasts stay in the same place, cp-actin filaments were few, but when chloroplast-avoidance movement was induced, thick cp-actin filaments appeared on the front part of chloroplasts, with almost none on the rear part. The speed of the chloroplast-avoidance response was dependent on the difference in cp-actin amounts between the front and rear faces of the chloroplasts; the bigger the difference, the faster the movement [136,137]. In an A. capillus-veneris gametophyte cell, a similar structure of actin filaments was observed on the front part of moving chloroplasts [138]. The chloroplast accumulation response was induced by microbeam irradiation at the central part of a dark-adapted gametophyte cell. When the accumulation response became obvious, the gametophyte was pre-fixed with m-maleimidobenzoic acid N-hydroxysuccinimide ester (MBS), then fixed with formaldehyde, and finally stained overnight with Alexa Fluor 488-labeled phalloidin. The actin filaments were observed at the front part of the chloroplasts [138]. Actin ring structures surrounding the chloroplasts were also observed in protonema cells when the accumulation response was induced. These structures disappeared before the chloroplasts moved away from the accumulation area, indicating that these structures might not be involved in the induction of the chloroplast movement but that they may anchor the chloroplasts to the plasma membrane [60]. Protein factors involved in chloroplast movement Several mutant lines deficient in chloroplast movement have been isolated in A. capillus-veneris. Screening of mutant lines with distinguishable phenotypes is quite straightforward in fern gametophytes, as explained above, but subsequent identification of the mutated gene in ferns is very difficult. On the other hand, many chloroplast movement mutants have been isolated in A. thaliana, and the genes involved were subsequently identified. These include non-phototropic hypocotyl-like 1 (NPL1, [118]), renamed PHOT2, chloroplast unusual positioning 1 (CHUP1, [139]), J-domain protein required for chloroplast accumulation response 1 (JAC1, [140]), plastid movement impaired 1 (PMI1, [141]), PMI2 [142], weak chloroplast movement under blue light 1 (WEB1, [143]), kinesin-like protein for actin-based chloroplast movement 1 (KAC1, [144]) and thrumin1 (THRUM1, [145]). Genomic analyses of seed and lower plants have revealed that CHUP1, KAC and PMI1 are also found in lower plants but THRUM1, JAC1, PMI2 and WEB1 are specific to seed plants [146]. In chup1 and kac1 mutants of Arabidopsis, no cp-actin filaments were observed, suggesting involvement of these genes in actin polymerization or actin filament maintenance [136,144]. Based on phenotypes similar to those of the characterized Arabidopsis mutants phot2 [38], chup1 and kac1 [147], mutants were identified in A. capillus-veneris, meaning that fundamental factors for cp-actin polymerization or maintenance are widely distributed in all plants, and that the basic mechanisms of chloroplast movement using cp-actin filaments are conserved [146]. In addition, phy3 mutant lines were identified in A. capillus-veneris as a-phototropic mutants under red light [113]. phy3 was also the photoreceptor for chloroplast movement [17], and was renamed neochrome (neo), as mentioned earlier [65]. Nuclear photorelocation movement Similarly to chloroplasts, nuclei also show photorelocation movement in A. capillus-veneris as well as in A. thaliana [148–151], for reviews, see [152,153]. This phenomenon was first described by Senn in his book published in 1908. Recently, it was reported that nuclear movement towards the anticlinal walls in response to strong light is effective for DNA damage protection in A. thaliana [154]. In A. capillus-veneris prothallial cells growing under low light conditions, be it white, red, or blue, the nuclei are in the center of the cells, but the nuclei cannot be seen without staining because they are under chloroplasts, which spread over the periclinal wall [148]. In the dark, the nuclei move to the anticlinal wall together with the chloroplasts [148]. Nuclei showed an accumulation response under horizontal irradiation (parallel to the prothallial surface) with polarized red or blue light, moving to the walls parallel to the vibration plane of the polarized light [149]. Under high light conditions with white or blue light irradiated vertically (perpendicular to the prothallus), nuclei at the periclinal wall moved to the anticlinal wall together with chloroplasts [150]. Experiments using phot2 and neo1 mutants showed that the photoreceptor for the avoidance response was phot2, which also mediated the dark-positioning of the nuclei, while the red-light-induced nuclear accumulation movement was mediated by neo1. The blue-light-dependent accumulation movement of the nuclei was redundantly mediated by phot2 and neo1, and probably by phot1 [150]. The mechanism of light-induced movement of nuclei in ferns is not known; however, in Arabidopsis mesophyll and pavement cells, light-induced nuclear movement is carried out by plastids and the nuclei cannot move by themselves [155]. Similar mechanisms might function in prothallial cells because the nuclei are suspended under chloroplasts in prothallial cells [148]. Concluding remarks In protonemal cells of the fern A. capillus-veneris, elementary processes of plant development, such as cell growth, phototropic response, cell division, timing of the cell cycle and direction of cell division, are controlled at each cellular stage by red- or blue-light irradiation. Hence, it is possible to make long protonemal cells of the desired length and shape, such as L-shaped, S-shaped, even branched cells [26]. Two-dimensional prothallial cells are also useful as nonpolar cells with or without chloroplasts on their periclinal walls [42]. The gametophytes are not surrounded with any tissues, so that light irradiation, cell manipulation, and observation of the results are easy. Using these advantages of fern gametophytes, we have been able to clarify cell biological processes in addition to photobiological ones, such as the involvement of the nucleus on the position of PPB formation and in its disappearance [75,80,84], and the measurement of the transmission speed of the signal for the chloroplast accumulation response [100,133]. Branched protonemal cells with two apices grow at either one if they are filled with cytoplasm and a nucleus by cell centrifugation [91]. This system might be useful for the study of polarity. Although there is great potential for the use of fern gametophytes as tools for the analysis of complex physiological processes, little attention is being paid to them because stable gene transformation of gametophytes is almost impossible at the moment. However, gene transfer has been shown to be possible by particle bombardment [17,38], although the effect is transient. Moreover, DNAi (DNA interference, similar to RNA interference) is a useful technique to silence the gene with a sequence, and the effect is not restricted in the bombarded cell but spreads to the entire cells of the gametophyte [156]. We hope that genetic transformation protocols suitable for fern gametophytes may be developed in the near future, so that the tremendous potential of the fern gametophyte system will be finally realized. Acknowledgments I would like to thank Prof. Yoshinobu Mineyuki inviting me to write this review and Prof. Satoru Tokutomi who kindly drew Fig. 1 and Dr Eiji Gotoh for arranging figures. I also thank my former colleagues supporting the works described in this review, especially Dr Hidenori Tsuboi for his valuable suggestions and Dr Takeshi Higa for careful reading of this manuscript. 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TI - Light-dependent spatiotemporal control of plant cell development and organelle movement in fern gametophytes
JO - Microscopy
DO - 10.1093/jmicro/dfy143
DA - 2019-02-01
UR - https://www.deepdyve.com/lp/oxford-university-press/light-dependent-spatiotemporal-control-of-plant-cell-development-and-ArRvFsZSyd
SP - 13
VL - 68
IS - 1
DP - DeepDyve
ER -