Ulva mutabilis Føyn has attracted the interest of phycologists since its first isolation by Føyn (1958), who established this species as a laboratory organism in 1952. Many studies were published during the 1960s and 1970s mainly on the ultrastructure of the thallus, cell division and morphogenesis (Løvlie 1964, 1968, Løvlie and Bråten 1968, 1970, Hoxmark and Nordby 1974, Fjeld and Løvlie 1976).
A detailed description of the thallus of the developmental mutant “slender”, which is often used for experimental studies because of its shorter developmental cycle, was given first by Løvlie and colleagues (Løvlie 1964, 1968, Fjeld and Løvlie 1976) and has been further analysed and confirmed by Wichard and Oertel (2010). An important analysis of the vegetative cell cycle by radioactive labelling with 14C-uracil confirmed that the cell cycles were synchronous and governed by a circadian rhythm (Stratmann et al. 1996).
Regarding cultivation, sexual reproduction represents a fundamental phase linked to the production of gametes. Blade cells of U. mutabilis excrete regulatory factors into their cell walls and the environment. Upon removal of the sporulation inhibitors, the gametogenesis of the thalli can be artificially and synchronously initiated (Stratmann et al. 1996, Wichard and Oertel 2010). This was also observed in Ulva linza L. (Vesty et al. 2015) and Ulva lactuca L. (Wichard and Oertel 2010).
As fragmentation of the thalli washes out the regulatory sporulation inhibitors of gametogenesis and zoosporogenesis, it triggered the propagation of Ulva (Stratmann et al. 1996) and this was confirmed by observation of the proliferation of Ulva prolifera O.F. Mueller during green tides (Gao et al. 2010).
Significant cell-biological changes have already been observed during gametogenesis using scanning electron microscopy (SEM) (Bråten 1971, Wichard and Oertel 2010) and, more recently, metabolic changes have been observed by matrix assisted laser desorption/ionisation mass spectrometric imaging (R.W. Kessler, A.C. Crecelius, U.S. Schubert, and T. Wichard, unpublished). Around noon of the second day after induction of gametogenesis, when the blade cells were irreversibly committed to the differentiation of gametangia, the chloroplasts appeared reoriented, and papillae started growing at the outer surface of the blade cells (Figure 1). The papillae converted, finally, into “capped” pores, while all progametes matured into 16 fully differentiated biflagellate gametes overnight. In the morning, the “caps” were successively dropped and the open exit pores became visible. Although most pores seemed to be open at this time and the gametangia were exposed to light, the mature gametes remained completely motionless because of the presence of a swarming inhibitor (SWI) in the medium. After removal of the SWI, the gametangia were completely discharged within a few minutes (Wichard and Oertel 2010, Vesty et al. 2015).
Although there are a large number of publications on the development, genetics and physiology of U. mutabilis, the cell structure and the cytoskeleton organisation during gametogenesis have not yet been studied in detail. The only available information derives from the original ultrastructural studies of fertilisation and zygote formation (Bråten 1971), of meiosis and centriole behaviour (Bråten and Nordby 1973), and from light microscopy of zoosporogenesis and gametogenesis of Ulva (Nordby 1974). Therefore, gametogenesis of Ulva is a very intriguing process for understanding the function and dynamic changes of the cytoskeleton and its role in the mechanism of gametangia formation and the gamete release.
The main aim of the present study was, therefore, to elucidate the mechanism of gametogenesis by examining the fine structural changes occurring during this process with particular attention to the cell wall differentiation and the microtubule cytoskeleton organisation under standardised culture conditions.
Materials and methods
Algal material and cultivation
Haploid gametophytes from the fast-growing developmental mutant “slender” (sl) of Ulva mutabilis Føyn (mating type mt+) were cultured. The sl-G(mt+) mutant used was a derivative of the original sl strain, selected in the laboratory, which had lost its capability for spontaneous diploidization during parthenogenetic propagation (Hoxmark 1975, Løvlie and Bryhni 1978). Ulva mutabilis was cultivated in Ulva culture medium (Stratmann et al. 1996) in a 17:7 h light/dark regime at 18°C with an illumination of 80–120 μmol photons m−2 s−1 (50% GroLux, 50% daylight fluorescent tubes; OSRAM, München, Germany) and no additional aeration. Further information about the materials used, the culture conditions and the preparation of the seed stock can be found in Wichard and Oertel (2010). All chemicals were purchased from Sigma-Aldrich (Taufkirchen, Germany) or Merck (Darmstadt, Germany).
Induction of gametogenesis
Intact mature thalli of the naturally occurring developmental mutant “slender” of Ulva mutabilis were washed for 15 min with distilled water at noon and minced manually into 3–5 mm fragments using an herb chopper (Zyliss, Zürich, Switzerland) prior to washing and distribution in fresh medium. The gametangia were mature in the morning of the third day after this medium change, and swarming of the gametes was induced by an additional medium change and application of light. Thallus fragments were collected directly before and every 24 h after, induction of gametogenesis for tubulin immunofluorescence staining, TEM and light microscopy. The fragments were inspected under a microscope to determine the status of gametogenesis (Figure 1). The following protocols were used.
The indirect immunofluorescence method was applied for the localization of tubulin, following a modified protocol described by Katsaros and Galatis (1992), i.e. small thallus pieces were fixed in 4% paraformaldehyde (PFA) and 0.125% glutaraldehyde in microtubule stabilising buffer (MTB; v:v; 50 mmol l−1 PIPES, 5 mmol l−1 ethyleneglycol-bis(aminoethyl ether)-tetra-acetic acid: EGTA, 5 mmol l−1 MgSO4·7H2O, 25 mmol l−1 KCl, 4% NaCl, 2.5% polyvinyl pyrrolidone: PVP 25, v:v, 1 mmol l−1 DL-dithiothreitol: DTT, pH 7.4) for 1 h. The samples were then washed thoroughly with MTB and the latter was gradually replaced by phosphate-buffered saline (PBS; 137 mmol l−1 NaCl, 0.7 mmol l−1 KCl, 5.1 mmol l−1 Na2HPO4, 1.7 mmol l−1 KH2PO4, 0.01% NaN2, pH 7.4, v:v). After washing, the samples were transferred to the enzyme digestion solution, i.e. PBS containing 2% cellulysin (219466, Lot No. B19227, Calbiochem-Novabiochem Corporation, San Diego, CA, USA), 5% hemicellulase (H2125-150KU, Lot. No SLBH6404V, Sigma-Aldrich), 1% driselase (D-8037, Lot No. 19H0784, Sigma-Aldrich), 2% pectinase (P-9179, Lot No. 53H0684, Sigma-Aldrich), 2% macerozyme (M8002.0001, Lot No. 010572.03, Duchefa Biochemie, Harrlem, The Netherlands) and 3% PMSF (a protease inhibitor). After washing with PBS, the thallus pieces were gently squashed on cover-slips covered with poly-L-lysin. The samples were left to dry and extraction of the pigments by Triton X-100 4% for 45 min at room temperature followed. After washing with PBS, incubation with the first antibody YOL 1/34 (Serotec, Puchheim, Germany), diluted 1:100 in PBS with 1%, (v:v) bovine serum albumin (BSA), continued overnight at room temperature. Washing again with PBS was followed by incubation with the secondary antibody anti-rat IgG FITC-conjugated (Sigma-Aldrich), diluted 1:100 in PBS-BSA for 2 h at 37°C in a dark and humid environment. The samples were then washed with PBS and treated for DNA staining with Hoechst 33258 (Molecular Probes, Eugene, OR, USA) in a final concentration of 5 μg ml−1 in PBS for 10 min at room temperature (Katsaros and Galatis 1992). Finally, the cover-slips were mounted on slides with a drop of anti-fade substance (0.0024 g p-phenylen-diamine, 1 ml glycerol and 0.5 ml PBS).
Transmission electron microscopy
In order to study the cell structure at different stages of gametogenesis, samples were taken at different times after the induction of gametogenesis, fixed primarily with 2% glutaraldehyde and 2% formalin (v:v; Merck) in 0.1 m cacodylate buffer (Sigma-Aldrich) and then post-fixed with 2% osmium tetroxide (v:v; Sigma-Aldrich). The samples then underwent electron microscopy according to established protocols (Løvlie and Bråten 1970).
The immunofluorescence samples were examined under epifluorescence with a Zeiss Axioplan™ microscope (Carl Zeiss, Oberkochen, Germany). Pictures were recorded with AxioCam Mrc 5 (Carl Zeiss) using the AxioVision™ software. In order to ensure the accuracy of the results, at least 50 cells with the same microtubule cytoskeleton were examined for each stage of gametogenesis. In parallel, gametogenesis was observed in living material by the same Zeiss Axioplan™ microscope described above or under a Leica DMIL LED microscope (Leica, Solms, Germany) equipped with a digital camera DS-Fi2 (Nikon, Düsseldorf, Germany). Thin sections were examined under a Philips EM 300 transmission electron microscope.
Interphase somatic cells
The thallus appeared in cross-section to consist of two cell layers which separated easily forming a monolayer (Figure 2). The cells were orthogonal in shape with the dimensions 16–30 μm length and 8–18 μm width (Figure 3). The undifferentiated cells of the developing thallus were characterised by a more or less central nucleus and a large cortical chloroplast with an obvious pyrenoid. The pyrenoid was traversed by a single thylakoid and surrounded by starch grains, while smaller starch grains were observed among the thylakoid membranes. The cell wall was thick and consisted of several layers (Figures 3 and 4): the internal one is the wall of each cell “itself”. This internal part consisted of a few thin parallel bundles of fibrillar material (probably cellulose microfibrils) embedded in a large amount of amorphous material. A second multilayered wall covered all the cells externally. This external wall part consists of one electron-dense external thin layer and a number of other internal layers, which bore thicker and electron-dense fibrillar bundles (Figure 4).
The microtubule cytoskeleton at this stage consisted of more or less parallel microtubule bundles arranged mainly in the cortical cytoplasm parallel to the plasmalemma (Figure 9).
Early stages of gametangial formation
The first sign of the transformation of somatic cells to gametangia was observed 48 h after the induction of gametogenesis by the formation of a slight projection (papilla) towards the outside of the thallus. The cells appeared polarised with most of the cytoplasm and the chloroplast located at the tip and the vacuoles at the base, while the nucleus was located in a more central position (Figure 5). The external fraction of the cell wall appeared to be differentiated.
The internal part of the cell wall pushed the layers above which seemed to be breaking apart (Figures 5 and 6). The parallel layers of cellulose microfibrils were interrupted and a round structure of amorphous material was formed (Figure 7). Changes in the chemical composition of the cell wall occurred at the same time, as an increase of the thin external dark-stained wall layer could be observed (Figures 6 and 7). The dark-stained material appeared to be extending gradually and covered all the round wall area, forming a “cap-like” structure (Figure 8).
The microtubule cytoskeleton was also drastically changed during this stage. The cortical microtubule bundles disappeared and a new system of intensely fluorescent cortical microtubule bundles appeared running parallel to the cell axis and converging towards the conical cell projection. Most of the microtubules were observed in the first stages of gametangial formation around the top of the conical projection (Figure 10). In more advanced stages, the microtubule cytoskeleton was further changed, forming thin bundles which radiated out of a particular point at the base of the cell (Figure 11), traversed the cell periphery in a basket-like shape and converged to a circular area behind the top of the conical cell projection (Figures 12 and 14). However, the microtubule bundles did not reach the top, leaving a circular opening between them (Figures 12 and 13).
Advanced stages of gametangial formation – gamete formation
After the completion of gametangia formation, repeated mitotic divisions gave rise to 16 gametes in each gametangium. These divisions were accompanied by a reorganisation of the cytoplasm, with the loss of its previous polar organisation. No centrosomes or any similar polar structures were found organising the spindle, and the nuclear envelope remained intact during the first mitotic stages, breaking only at the poles (Figures 15 and 16). The cortical microtubule system of the previous stage disappeared and the spindle was organised by microtubules which entered the nucleoplasm through the polar gaps. During anaphase-telophase, the spindle was elongated and microtubule bundles appeared in the interzonal area. Cytokinesis started parallel to the mitotic divisions by furrowing membranes which extended between the daughter nuclei (Figures 17 and 18). Increased numbers of dictyosomes and endoplasmic reticulum (ER) membranes were observed close to the developing furrowing membranes (Figure 18). It was interesting that the cytokinetic membranes separating neighbouring and almost fully developed gametes in mature gametangia were not completed, leaving a narrow cytoplasmic bridge connecting them (Figures 19 and 21). The membrane surrounding these cytoplasmic bridges was lined by a dark-stained material (Figures 22 and 23). The microtubules also traversed the cortical cytoplasm of this area. Parallel to the mitotic divisions, a pair of centrosome-like structures was formed de novo in each daughter cell. These structures were rod-shaped and consisted of microtubule doublets in a circular configuration. The dark-stained material was connected with the base of these structures which were usually not connected to the nuclei, but attached to the external membrane of the gametangium (Figures 15 and 17).
The newly formed gametes had a round shape, which later became spindle-like, and were equipped with a pair of flagella arising from a basal body (Figures 20 and 21). They were usually oriented with their long axes converging towards the conical aperture of the top. The microtubule cytoskeleton in the individual gametes consisted of cortical bundles radiating out of a particular point, which was the basal body of the flagellum.
At this stage, the cell wall of the tip region was further differentiated. The external layer of the cell wall appeared broken and the internal layer underwent a particular differentiation, with the deposition of additional layers of wall material forming a plug-shaped cap, closing the opening of the external wall layers (Figure 24). The plug consisted of parallel layers of cellulose microfibrils showing a rather loose pattern (Figure 25). Interestingly, at this stage, a dark-stained amorphous material was visible under the opened exit pores (Figure 26).
After the completion of the flagella formation, the plug-shaped cap was removed and gamete discharge could take place through the opening of the release tube. Anti-tubulin immunofluorescence staining revealed that each gamete bore two flagella (Figures 27–30). The microtubule cytoskeleton of the gametes released consisted of thick, intensely fluorescent bundles surrounding the cell cortex (Figure 27). Apart from the cortical microtubules, thinner and shorter bundles and possibly single microtubules traversed the cytoplasm in other directions (Figures 29 and 30). All the microtubules of the gametes radiated out of the flagellar basal body region.
The process of gametangia formation and gamete release in Ulva mutabilis is highly synchronised. A crucial role in this process is played by the presence of the sporulation and swarming inhibitors (Stratmann et al. 1996, Wichard and Oertel 2010). By removal of the sporulation inhibitors gametogenesis can be artificially induced in the laboratory. A particular divergence in the differentiation is followed in order to transform somatic cells to gametangia. This process includes drastic changes in the cell structure. The polarisation of the cell, which is expressed by the rearrangement of the cell elements, is accompanied and probably controlled by the transformation of the microtubule cytoskeleton. It has been reported that interphase cells of Ulva lactuca bear an impressive microtubule system of intensely fluorescing microtubule bundles which radiate from a cortical site close to the nucleus and surround the chloroplast (Tsagkamilis and Katsaros 2007). However, it was not clear in this study whether these cells were at a preliminary gametogenesis stage. From both the old preliminary and the current findings, it is clear that the microtubule organisation in interphase cells of Ulva is different from land plants (Hashimoto 2015 and literature therein), therefore additional work is needed to clarify the role of microtubules in cell wall morphogenesis. The conical microtubule organisation may facilitate the transport of cell elements to the top of the cell. A similar role of the microtubule cytoskeleton has been reported in other polarised systems, such as in chlorenchyma cells of flowering plants (Chuong et al. 2006), bryophytes (Doonan et al. 1988), fungal hyphae (Roberson and Vargas 1994), green algae (Holzinger et al. 2002) and brown algae (Karyophyllis et al. 1997, Katsaros et al. 2006). However, actin has also been reported to participate in organelle movement (Grolig 1990, Higaki et al. 2007, Blanchoin et al. 2010) . Therefore, it would be interesting to examine the role of actin in the organisation of the cytoplasm of the developing gametangia in future studies.
The second interesting process occurring during gametogenesis is the differentiation of the cell wall. As has already been mentioned, the cell wall of the gametangium consists of two electron-dense layers: an outer layer representing the vegetative cell wall and an inner layer or capsule. This structure has been described in sporangia of U. mutabilis (Løvlie and Bråten 1968) and in gametangia of U. intestinalis L. (McArthur and Moss 1979), as well as in the sporangial wall of Endophyton ramosum N.L. Gardner (Leonardi et al. 1997). A more detailed description of the cell wall of Ulva compressa L. has been reported by Holzinger et al. (2015). According to this study, the wall is multilayered, consisting of cellulose, callose and acidic polysaccharides. This architecture provides particular desiccation tolerance in Ulva. The capsular zoosporangial wall and method of sporangial dehiscence (i.e. wall dissociation and the production of a mucilage plug) were characters of the Ulvales sensu Stewart and Mattox (1975).
The differentiation of the external cell wall finally results in the formation of the exit pore for the release of gametes. Similar observations have been reported during sporangia formation and meiotic divisions of U. mutabilis sporophytes (Bråten and Nordby 1973). In this study, only the final stage before the opening was presented showing less densely packed and more randomly oriented fibrils than in the rest of the cell wall. This was an indication of the cell wall weakening before the opening of the exit pore. However, we found that the process of cell wall differentiation started very early, even 36 h after the induction of gametogenesis. The first sign was the change of the chemical composition of the cell wall. It seems that the induction of gametogenesis triggered the activation of a mechanism of synthesis of new, rather amorphous wall material, which was guided to the top of the cell. This material was concentrated gradually in a limited area, interrupting the continuity of the cell wall layers. Bråten and Nordby (1973) found membrane-bound vesicles at the site of cell wall weakening. Such vesicles could probably transfer cell wall precursor materials which contributed to the bulging. Wichard and Oertel (2010) reported that wall papillae start growing at the outer surface of the cells when they were already irreversibly committed to gametangium differentiation, the chloroplasts appeared reoriented and the first progametes had divided. The papillae converted into “capped” pores during the following night (Figure 1). Using TEM, we found that the process of “cap” formation started quite early by the gradual differentiation of the cell wall.
A third important finding of the present study was the reorganisation of the microtubule cytoskeleton during the gametangia formation and gamete release. The cortical, basket-like microtubule configuration showed two focal sites. The site at the bottom (internal side) of the cell is possibly functioning as a microtubule organising centre (MTOC). Since we did not find a centrosome in the interphase cells of U. mutabilis that would organise microtubules, we can assume that interphase microtubules are organised by proteins which do not form a visible structure, similar to the interphase of land plant cells, and this supports the phylogenetic relationship between green algae and higher plants (Meagher and Williamson 1994 , Wasteneys 2004). However, the fact that the nuclear envelope does not disassemble during mitosis, and the spindle microtubules enter the nucleoplasm through the polar gaps is characteristic of lower eukaryotes and algae rather than of land plants (Güttinger et al. 2009, Boettcher and Barral 2013). As noted in the results, these microtubules converged to a circular area at the top of the cell. This fits very well with the architecture of the gametangium, meaning that microtubules reached the “neck” of the exit pore. From this particular organisation, it can be reasonably suggested that microtubules are implicated in the gametangia maturation.
In the following stages, repeated nuclear divisions occurred to form the 16 gametes. The absence of centrosome-dependent MTOCs in the spindle poles of these mitoses suggests that the spindle was also organised without the involvement of the centrosomes. However, centrosome-like structures were formed parallel to the divisions. These structures were close to the nuclei, and were not connected with them, but were attached to the developing furrowing membrane. A similar process has been described by Bråten and Nordby (1973) during the sporogenesis of U. mutabilis.
An interesting phenomenon observed was the cytokinetic process during the gamete formation, i.e. the mechanism of separation of individual gametes. Since the ready-to-release gametes were surrounded only by a membrane and not a cell wall, it was expected that a cytokinetic diaphragm would be developed between each pair of daughter nuclei. This process started quite early in the present study, after telophase, with the furrowing of the plasma membrane (Figure 17). This mechanism differs from that described in other multinucleate systems, such as in endosperms of flowering plants, where cytokinesis took place after the complete formation of the daughter nuclei (Sørensen et al. 2002). The presence of an increased number of active dictyosomes and membranes of the endoplasmic reticulum (ER) close to the developing furrow suggests that vesicles from golgi and ER may have contributed to the development of this separating diaphragm.
Interestingly, cytoplasmic connexions were observed in almost fully developed gametes constituting a kind of network by which all the gametes were connected within the gametangium. These cytoplasmic bridges are similar to the connexions between the fusing gametes of U. mutabilis observed by Bråten (1971). The cytoplasmic bridges in Ulva may also be compared with those described in Volvox L. embryos by Green et al. (1981). They are formed as a result of incomplete cytokinesis and it has been suggested that they play a role in generating the movements of the inversion process of Volvox. Therefore, we hypothesise an important role for the cytoplasmic bridges in the gamete release mechanism of Ulva. This could also be combined with the period when the gametes are motionless due to the presence of the SWI as well as with the fact that treatment with the non-competitive myosin-ATPase blocker 2,3-butane-dione monoxime (BDM) caused complete inhibition of gamete release (Wichard and Oertel 2010). The dark-stained material underlining the cytoplasmic channel could be an actin ring. This ring may contribute to the final closing of the gap and the separation of gametes, thus, facilitating their synchronous release one after the other.
After the complete formation of the gametes, their release through the opened exit pore occurred. The microtubule cytoskeleton of each gamete consisted of cortical bundles diverging from the basal body site. Two flagella rose from the basal body. An interesting observation during the gamete release was that the oval-shaped gametes showed a kind of orientation with their long axis converging towards the opening. This means that a specific mechanism underlies this process. Experiments with the myosin-ATPase blocker BDM have indicated that the mechanism of gamete release depends potentially on a myosin-ATPase. Although the inhibiting concentration of 40 mmol l−1 reported was high (Wichard and Oertel 2010), it is similar to the inhibiting concentration found in diatoms, where gliding is based on an actin/myosin system (Poulsen et al. 1999).
The process of gamete release may involve a mechanism by which the gametes are mechanically extruded out of the gametangium. This may occur by hydration of an extracellular substance (possibly callose-like) which causes a considerable increase of its volume and, in turn, pushes the gametes out of the gametangium. Leonardi et al. (1997) studied the development and morphology of sporangia and zoospores of the filamentous Ulvophyte Endophyton. They found that the sporangium was filled with a mucilagenous fibrillar material in which mature zoospores remained immersed until liberation. A thick layer of similar material covered the inner sporangial cell wall and the upper apical part of the sporangium. Our observations of dark-stained material below the exit pore of U. mutabilis gametangia (Figure 26) supported this hypothesis. In this context, the fragile inner perforated seal reported by Wichard and Oertel (2010) could not be observed by TEM.
As all the above data have been extracted from the naturally occurring developmental mutant “slender”, future work should also include comparison with the wild type alga. However, differences in regulatory effects, such as sporulation and swarming inhibitors, between morphotypes or even species have not been observed up to now (Wichard and Oertel 2010, Vesty et al. 2015).
Interestingly, under axenic conditions, Ulva forms callus-like cultures of blade cells which do not undergo gametogenesis (Spoerner et al. 2012, Wichard 2015). However, comparative studies of the microtubule organisation of axenic cultures with Ulva cultures associated with the natural microbiome will give further detailed evidence on how microtubules contribute to bacteria-induced cell differentiation processes.
While certain cytoskeletal features are widely conserved across different plant phyla, including the highly developed cortical cytoskeleton domains associated with developmental processes, each macroalgal species possesses its own unique specialisation. The synchronised formation of the exit pore is certainly crucial for the mating success of gametes. Simultaneous formation of both gametes and the exit pores was observed, whereas the discharge of gametes can be synchronised by the SWI which increases the mating probability of the two different mating types. The methodology described here not only paves the way for monitoring the microtubule-based trafficking during gametogenesis, but also, for example, for the photoreceptor and eyespot localization which is an essential process for the phototaxis of germ cells of Ulva. Using the newly developed transformation system in Ulva (Oertel et al. 2015), the structure and functioning of the cytoskeleton controlled by associated proteins (Timmers et al. 2002) can be investigated for the first time, for example, by green fluorescent fusion proteins in macroalgae. Even more interestingly, as axenic Ulva cultures develop into a callus-like morphotype (Spoerner et al. 2012, Wichard 2015), our further research will explore how the bacterial morphogenetic substances interfere with the microtubule network in cell organisation, function and morphogenesis.
This study was financed by the National and Kapodistrian University of Athens (“Kapodistrias” programme) to C.K and by the Deutsche Forschungsgemeinschaft (DFG) through the Jena School for Microbial Communication (German Excellence Initiative) and the Collaborative Research Center SFB 1127 “ChemBioSys” to A.W. and T.W. The authors would like to acknowledge networking support by the European Cooperation in Science and Technology (COST) Action “Phycomorph” FA1406.
Blanchoin, L., R. Boujemaa-Paterski, J.L. Henty, P. Khurana and C.J. Staiger. 2010. Actin dynamics in plant cells: a team effort from multiple proteins orchestrates this very fast-paced game. Curr. Opin. Plant Biol. 13: 714–723.
Boettcher, B. and Y. Barral. 2013. The cell biology of open and closed mitosis. Nucleus. 4: 160–165.
Bråten, T. 1971. Ultrastructure of fertilization and zygote formation in green alga Ulva mutabilis Føyn. J. Cell Sci. 9: 621–635.
Bråten, T. and O. Nordby. 1973. Ultrastructure of meiosis and centriole behavior in Ulva mutabilis Føyn. J. Cell Sci. 13: 69–81.
Chuong, S.D., V.R. Franceschi and G.E. Edwards. 2006. The cytoskeleton maintains organelle partitioning required for single-cell C4 photosynthesis in Chenopodiaceae species. Plant Cell. 18: 2207–2223.
Doonan, J., D. Cove and C. Lloyd. 1988. Microtubules and microfilaments in tip growth: evidence that microtubules impose polarity on protonemal growth in Physcomitrella patens. J. Cell Sci. 89: 533–540.
Fjeld, A. and A. Løvlie. 1976. Genetics of multicellular marine algae. In: (R.A. Lewin, ed) The genetics of algae. Botanical monographs. Univ. Calif. Press, Berkeley, CA. pp. 219–235.
Føyn, B. 1958. Über die Sexualität und den Generationswechsel von Ulva mutabilis. Arch. Protistenkd. 102: 473–480.
Gao, S., X.Y. Chen, Q.Q. Yi, G.C. Wang, G.H. Pan, A.P. Lin and G. Peng. 2010. A strategy for the proliferation of Ulva prolifera, main causative species of green tides, with formation of sporangia by fragmentation. Plos One. 5: e8571.
Green, K.J., G.I. Viamontes, and D.L. Kirk. 1981. Mechanism of formation, ultrastructure, and function of the cytoplasmic bridge system during morphogenesis in Volvox. J. Cell Biol. 91: 756–769.
Grolig, F. 1990. Actin-based organelle movements in interphase Spirogyra. Protoplasma. 155: 29–42.
Güttinger, S., E. Laurell and U. Kutay. 2009. Orchestrating nuclear envelope disassembly and reassembly during mitosis. Nat. Rev. Mol. Cell Biol. 10: 178–191.
Hashimoto, T. 2015. Microtubules in plants in The Arabidopsis Book: American Society of Plant Biologists 13: e0179.
Higaki, T., T. Sano and S. Hasezawa. 2007. Actin microfilament dynamics and actin side-binding proteins in plants. Curr. Opin. Plant Biol. 10: 549–556.
Holzinger, A., S. Monajembashi, K. Greulich and U. Lütz-Meindl. 2002. Impairment of cytoskeleton-dependent vesicle and organelle translocation in green algae: combined use of a microfocused infrared laser as microbeam and optical tweezers. J. Microsc. 208: 77–83.
Holzinger, A., K. Herburger, F. Kaplan and L.A. Lewis. 2015. Desiccation tolerance in the chlorophyte green alga Ulva compressa: does cell wall architecture contribute to ecological success? Planta. 242: 477–492.
Hoxmark, R.C. 1975. Experimental analysis of life cycle of Ulva mutabilis. Bot. Mar. 18: 123–129.
Hoxmark, R.C. and O. Nordby. 1974. Haploid meiosis as a regular phenomenon in life cycle of Ulva mutabilis. Hereditas. 76: 239–249.
Karyophyllis, D., B. Galatis and C. Katsaros. 1997. Centrosome and microtubule dynamics in apical cells of Sphacelaria rigidula (Phaeophyceae) treated with nocodazole. Protoplasma. 199: 161–172.
Katsaros, C. and B. Galatis. 1992. Immunofluorescence and electron-microscopic studies of microtubule organization during the cell-cycle of Dictyota dichotoma (Phaeophyta, Dictyotales). Protoplasma. 169: 75–84.
Katsaros, C., D. Karyophyllis and B. Galatis. 2006. Cytoskeleton and morphogenesis in brown algae. Ann. Bot. 97: 679–693.
Leonardi, P.I., J.A. Correa and E.J. Cáceres. 1997. Ultrastructure and taxonomy of the genus Endophyton (Ulvales, Ulvophyceae). Eur. J. Phycol. 32: 175–183.
Løvlie, A. 1964. Genetic control of division rate and morphogenesis in Ulva mutabilis Føyn. C R Trav Lab Carlsberg. 34: 77–168.
Løvlie, A. 1968. On the use of a multicellular alga (Ulva mutabilis Føyn) in the study of general aspects of growth and differentiation. Nytt. Magasin. for Zoologi. 16: 39–49.
Løvlie, A. and T. Bråten. 1968. On division of cytoplasm and chloroplast in the multicellular green alga Ulva mutabilis Føyn. Exp. Cell Res. 51: 211–220.
Løvlie, A. and T. Bråten. 1970. On mitosis in multicellular alga Ulva mutabilis Føyn. J. Cell Sci. 6: 109–128.
Løvlie, A. and E. Bryhni. 1978. On the relation between sexual an parthenogenetic reproduction in haplo-diplontic algae. Bot. Mar. 21: 155–164
McArthur, D. and B. Moss. 1979. Gametogenesis and gamete structure of Enteromorpha intestinalis (L.) Link. Br. Phycol. J. 14: 43–57.
Meagher, R.B. and R.E. Williamson. 1994. 38 The plant cytoskeleton. Cold Spring Harbor Monogr. Arch. 27: 1049–1084.
Nordby, O. 1974. Light microscopy of meiotic zoosporogenesis and mitotic gametogenesis in Ulva mutabilis Føyn. J. Cell Sci. 15: 443–455.
Oertel, W., T. Wichard and A. Weissgerber. 2015. Transformation of Ulva mutabilis (Chlorophyta) by vector plasmids integrating into the genome. J. Phycol. 51: 963–979.
Poulsen, N.C., I. Spector, T.P. Spurck, T.F. Schultz and R. Wetherbee. 1999. Diatom gliding is the result of an actin-myosin motility system. Cell Motil. Cytoskeleton. 44: 23–33.
Roberson, R. and M. Vargas. 1994. The tubulin cytoskeleton and its sites of nucleation in hyphal tips of Allomyces macrogynus. Protoplasma. 182: 19–31.
Sørensen, M.B., U. Mayer, W. Lukowitz, H. Robert, P. Chambrier, G. Jürgens, C. Somerville, L. Lepiniec and F. Berger. 2002. Cellularisation in the endosperm of Arabidopsis thaliana is coupled to mitosis and shares multiple components with cytokinesis. Development. 129: 5567–5576.
Spoerner, M., T. Wichard, T. Bachhuber, J. Stratmann and W. Oertel. 2012. Growth and thallus morphogenesis of Ulva mutabilis (Chlorophyta) depends on a combination of two bacterial species excreting regulatory factors. J. Phycol. 48: 1433–1447.
Stewart, K.D. and K.R. Mattox. 1975. Comparative cytology, evolution and classification of the green algae with some consideration of the origin of other organisms with chlorophylls a and b. Bot. Rev. 41: 104–135.
Stratmann, J., G. Paputsoglu and W. Oertel. 1996. Differentiation of Ulva mutabilis (Chlorophyta) gametangia and gamete release are controlled by extracellular inhibitors. J. Phycol. 32: 1009–1021.
Timmers, J.A.C., A. Niebel, C. Balagué and A. Dagkesamanskaya. 2002. Differential localisation of GFP fusions to cytoskeleton-binding proteins in animal, plant, and yeast cells. Protoplasma. 220: 69–78.
Tsagkamilis, P. and C. Katsaros. 2007. Microtubule organization during the cell cycle of the green alga Ulva lactuca. Proc. 29th Conf. of Hellenic Association for Biological Sciences, pp. 386–387.
Vesty, E.F., R.W. Kessler, T. Wichard and J.C. Coates. 2015. Regulation of gametogenesis and zoosporogenesis in Ulva linza (Chlorophyta): comparison with Ulva mutabilis and potential for laboratory culture. Front. Plant Sci. 6: 15.
Wasteneys, G.O. 2004. Progress in understanding the role of microtubules in plant cells. Curr. Opin. Plant Biol. 7: 651–660.
Wichard, T. 2015. Exploring bacteria-induced growth and morphogenesis in the green macroalga order Ulvales (Chlorophyta). Front. Plant Sci. 6: 86.
Wichard, T. and W. Oertel. 2010. Gametogenesis and gamete release of Ulva mutabilis and Ulva lactuca (Chlorophyta): regulatory effects and chemical characterization of the “swarming inhibitor”. J. Phycol. 46: 248–259.