Introduction

Formation of gametangia and fertilisation of charophytes

Sexual reproduction

The fusion of male and female gametes and the subsequent maturation of the zygote is called sexual reproduction. This mode of reproduction is described for mostly all charophyte species, even if the frequency of gametangia and oospore formation varies as can be seen in the enormous amount of available literature. Both gametangia, the male antheridia and the female oogonia were produced by the formation of short shoots from the nodal cell of branchlets (Schubert et al. 2016, Schubert et al. In press). Over the last 150 years, various authors published descriptions of cell development and formation of both gametangia. Due to the amount of considered aspects, not all of them can be considered here in detail. More than that, the plethora of techniques for gametangial studies range from simple descriptions over schematic drawings to high-resolution figures of microtome sections. Table 1 summarises available microtomic/microscopic studies of antheridia, oogonia and rhizoids. The full plethora of studies on internodal cells were excluded (Sundaralingam 1954, Fetzmann 1957, Barton 1965a, Barton 1965b, Nagai and Rebhuhn 1966, Pickett-Heaps 1966, Shen 1967c, Barton 1968, Vouilloud et al. 2015).

Male gametangium

The majority of detailed gametangial studies have dealt with male spermatozoids and have considered the morphology, cell division or phylogenetic relationship of charophyte spermatozoids to bryophytes or pteridophytes (Belajeff 1894, Mottier 1904, Karling 1926, Karling 1927, Sasaki 1935a, Sasaki 1935b, Satô 1954, Eggmann 1966, Pickett-Heaps 1967a, Pickett-Heaps 1967b). However, almost all comparisons are based on similarities in structure and form. In contrast to numerous evolutionary studies on the genetic regulations of male and female traits in bryophytes or pteridophytes, only a few studies are available for charophytes, such as the presence of the MIKC-type MADS box gene in oogonia and antheridia of Chara globularis (CgMADS1), suggesting the presence of these genes before diversification (Tanabe et al. 2004). These genes are thought to be involved in the control of sex-specific haploid cell differentiation and stress response (Qiu et al. 2023).

The development of the two flagellated spermatozoids is well known within this group of algae (Varley 1834, Thuret 1840). A brief historical overview of investigations is given by Vouilloud et al. (2012). In the 19^th century, different authors studied the cell development of all three cell types of antheridia (scutum, manubrium, capitulum) in detail (e.g. Varley (1834), Braun (1852), Pickett-Heaps (1967a)), a corresponding summary can be found in Schubert et al. (2016), Schubert et al. (In press). Intercellular streaming was only observed in the manubrium and capitulum, but never in external scutum cells (Braun 1852). In taxonomy, antheridia size is part of almost all species descriptions and has been used for species delimitation, for example for Tolypella species (Holzhausen et al. 2023). However, the developmental stage, environmental conditions, the reproduction mode and the position at the whorl can influence antheridia sizes (Ernst 1918, Kwiatkowska et al. 1996, Calero and Rodrigo 2022), which make it necessary to find a standardised method for reliable regional considerations or species delimitations by antheridia. Fig. 1 shows the typical development of antheridia which changed from green to light yellowish-green in younger stages to an orange antheridium in maturing stages, followed by the release of thousands of spermatozoids. In 1967, Pickett-Heaps suggested a correlation between the orange colour and the accumulation of plastoglobuli as a source of the increasing carotenoid content (Pickett-Heaps 1967a).

Figure 1.

Developmental stages of charophyte antheridia in accordance with Shepherd and Goodwin (1992). First developmental stage of young (A, B) and maturing (B) antheridia indicated by a green/light yellow colour or the orange colour (B) after shield cell developing (C) Autofluorescence of antheridia detected with a fluorescence microscope, scale = 139.8 µm. (D) Mature antheridia (E-F) release of spermatozoids by opening of shield cells (E) and a detailed view of antheridial filaments (F) (G) shield cells of antheridia after the release of spermatozoids containing plastids with plastoglobuli (photographs: A. Holzhausen).

In addition to morphology, there are several studies, mainly by Kwiatkowska and co-workers, on the cell-to-cell connection via plasmodesmata and the process of endoreplication in Chara vulgaris and/or Chara tomentosa (Kwiatkowska et al. 1990). The type of plasmodesmata (plugged, unplugged) correlates with the developmental stage and differs in the cell types connected and their internal substances (Kwiatkowska and Maszewski 1985, Kwiatkowska et al. 2003, Kwiatkowska 2003). Accordingly, Shepherd and Goodwin found out that the symplastic linkage in lateral branches between internodes or internodes and associated nodes is significantly higher in spring and summer (stage of harbouring young and maturing antheridia) than in the post-fertile stage (no new development of antheridia), the mature stage (antheridia reached at least a 32-celled-filament stage) or vegetative branches during winter (Shepherd and Goodwin 1992). In all these three stages, a restricted cell-to cell-communication was found by investigating the transport of 6COOHF in lateral branches. Broken plasmodesmata, therefore, determine the most mature state of antheridia and should be considered in further studies. Furthermore, autoradiographic studies have shown the circadian translational activity within the different cell types of antheridia (scutum, manubrium, capitulum, spermatozoids), including the protein synthesis activity of ³H-leucine (Kwiatkowska 1991, Kwiatkowska et al. 1995, Kwiatkowska et al. 1999, Kwiatkowska and Papiernik 1999). Studies on DNA methylation were performed by Olszewska et al. (1997)

Cell walls of Chara corallina antheridia show the presence of cellulose and pectin homogalacturon, whereas arabinogalactan proteins have only been indicated in epidermal shields and anti-xyloglucan in the capitulum (Domozych et al. 2009). The expression of PIN2-like auxin transporters in antheridial filaments was shown by immunofluorescence studies (Zabka et al. 2016). Studies on the effects of gibberellic acid are presented in subsection V, biomolecular studies.

Female gametangium and fertilisation

Compared to male antheridia, there are significantly fewer published studies on female unfertilised gametangia, so-called oogonia. While the male gametangium of charophytes (antheridia) is comparable to the antheridia of bryophytes, the two female gametangia differ. Charophyte oogonia are ovoid and the fertilisation leads directly to the zygote. In bryophytes, however, archegonia are multicellular and lageniform and fertilisation leads to sporophyte development, with the formation of spores (Braun 1852). Previous literature has concentrated on descriptions within general cryptogam overviews, on developmental stages of cell divisions by microtome sections and on size variations (e.g. De Bary 1871Covich and Tsukada (1969), Maier (1973), Sawa and Frame (1974), Leitch (1986), Leitch (1989), John et al. (1990), Leitch et al. (1990), Kwiatkowska et al. (1996)). The cell development of the female gametangium will not be discussed in detail here. Descriptions can be found in Braun (1852), Sundaralingam (1954), Schubert et al. (2016), Schubert et al. (In press). Microtome studies of female gametangia and included cell compounds are listed in Table 1. Intercellular streaming was observed only in the multicellular basal node, the unicellular internodal cell and the five envelope cells. However, the intercellular streaming within the oogonium could not be observed in detail (Braun 1852).

Table 1.

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List of cytological studies on gametangia, rhizoids and vegetative cell material of charophytes. The respective object and cell type, method, staining and identified organelles are recorded. If synonyms were used by the authors, common species name is listed first followed by the synonym in brackets.

Reference	Species	Method	Staining	Organelles
Johow (1881)	Chara vulgaris (=C. foetida) branchlet, gametangia, rhizoids, nodal cells	-	haematoxylin	nucleus division
Zacharias (1885), Zacharias (1888), Zacharias (1890)	C. vulgaris (=C. foetida) male and female gametangia, rhizoids	microscopy	-	nucleolus protein nuclei rhizoids geotropism
Overton (1890)	Nitella syncarpa, C. hispida female gametangium, branchlets and internodal cells	staining microscopy reaction against concentrated acids	chloralhydrate haematoxylin-eosin	nuclei of: stalk cells gametangial node cell “Wendezelle” spiral cells of oogonia coronula “Stachelkugeln”
Kaiser (1896)	C. vulgaris (=C. foetida), C. hispida, C. canescens (=C. crinita), N. syncarpa, N. flexilis apical cell, node cell, branchlets, oogonia/antheridia, cortex cells	sectioning (3-5 µm)	Alauncochenille Haidenhain’s Iron-haematoxylin haematoxylin-eosin in glycerine fuchsin methylene blue	karyokinesis nucleus nucleoli Altmann’s granula centrosomes
Goetz (1899)	N. flexilis, N. opaca, C. vulgaris (=C. foetida) internodal cells, oogonia/oospores	sectioning oogonia 10 µm sections zygotes 20 µm sections	fuchsin iodine green fuchsin methylene blue iron-haematoxylin maemalaun	chromatin nucleolus membrane
Mottier (1904)	C. globularis (=C. fragilis) spermatozoids	sectioning (3-5 µm)	aniline-safranin gentian-violet orange G.	nucleus cytoplasm blepharoplast cilia
Oelkers (1916)	C. globularis (=C. fragilis), C. vulgaris (=C. foetida), N. syncarpa generative/vegetative tissues	sectioning (30 µm) degradation of sugars	Haidenhain’s iron-haematoxylin	„Stachelkugeln“ Aleuron nuclei of zygotes
Riker (1921)	C. globularis (=C. fragilis), C. virgata (=C. verrucosa)	sectioning (5 – 7 µm)	Haidenhain’s iron-haematoxylin Flemming’s triple stain	mitochondria
Karling (1926)	C. braunii (=C. coronata), C. globularis (=C. fragilis), N. gracilis antheridia/oogonia, apical cells, nodal cells	microscopy	Heidenhain's iron-alum haematoxylin Flemming’s triple stain	shield cells manubria capitula filaments nuclei
Linsbauer (1927)	C. subspinosa (=C. rudis), C. vulgaris (=C. foetida), C. globularis (=C. fragilis) rhizoids	microscopy	methyl green acetic acid haematoxylin borax carmine	nucleolus rhizoidal cells
Schmuckler (1927)	C. contraria var. hispidula oogonia	microscopy	-	spiral cells of oogonia
Walther (1929)	N. syncarpa, N. hyalina, N. batrachosperma oogonia/antheridia, oospores, node cells, branchlets, internodal cells	sectioning (6-16 µm)	borax carmine acetic carmine methyl green Feulgen reagent	nuclei lipids starch
Sasaki (1935a), Sasaki (1935b)	N. sp., C. sp. spermatozoids	-	gentian-violet carbol-fuchsin	blepharoplast
Sundaralingam (1954)	C. zeylanica	sectioning (5-10 µm)	Haidenhain’s Iron-haematoxylin	antheridium oogonium oospores branchlets stem (cortex) internodal cells
Satô (1954)	C. braunii spermatozoids	electron microscopy phase contrast microscopy	carbol-fuchsin gentian-violet Heidenhain`s iron-haematoxylin Feulgen reaction aniline acidic fuchsin Janus green silver nitrate iodine iodide potassium solution	mitochondria chloroplast starch blepharoplast flagella flagella-fibrils spermatozoid-nucleus spherical bodies with granules
Barton (1965a), Barton (1965b)	C. vulgaris mature lateral cells	sectioning	-	plasmodesmata charasome mitochondria Golgi apparatus endoplasmic reticulum lipid bodies nucleus spherosomes
Hollenstein (1966)	C. sp. spermatozoids	X-ray diffraction	-	nuclein content
Shen (1967a), Shen (1967c)	C. contraria, C. zeylanica older internodes, rhizoids, protonema, leaves, shoot apices	sectioning (25 µm) smear whole plant part	fast Green Schiff’s reagent iron haematoxylin	nuclear DNA contents in mg sperm (22.5 ± 0.78) apical cell (45.7 ± 1.6) nodal cell (48.9 ± 1.59) nodal cell, metaphase (49 ± 1.23 mg) nodal cell, half-anaphase (21.9 ± 2.1 mg) 1-4 internode (124.2 ± 10.3 – 655.2 ± 45.3) crescentic nuclei, older internodes (174.2 ± 12.2) spherical nuclei (25.4 ± 1.3 / 46.6 ± 1.5)
Shen (1967b)	Chara zeylanica sperms	Feulgen spectral absorption	Schiff’s reagent	sperm nuclear DNA (9.46 ± 0.25 mg)
Pickett-Heaps (1967a), Pickett-Heaps (1967b), Pickett-Heaps (1968)	C. australis, C. fibrosa, C. sp., N. sp. antheridia oogonia	sectioning	uranium and lead toluidine blue	basal cell chloroplasts chromosomes endoplasmic reticulum cytoplasmic fibrils Golgi bodies nucleus mitotic spindle polysomes spermatogenous cell microtubules
Sawa and Frame (1974)	Tolypella nidifica plant apices with oogonia	sectioning (1-5 µm)	0.5% toluidine blue	sterile oogonial cell
Leitch (1986), Leitch (1989), Leitch et al. (1990)	C. aspera (=C. delicatula), C. hispida, Lamprothamnium papulosum oogonia	sectioning (80-100 nm; > 200 nm)	Reynold’s lead citrate 4% toluidine blue and 4% malachite green 4% basic fuchsin	basal position of nucleolus chemical composition and development of wall compounds/layers sphaerosomes
Moestrup (1970)	C. corallina spermatozoids	sectioning formvar/carbon-coated grids for electron microscopy	uranyl acetate Reynold’s lead citrate	different chromosome numbers in single antheridia filaments
Stabenau et al. (2003)	C. globularis (=C. fragilis) homogenates of all cells	sectioning	Reynold’s lead citrate and uranyl acetate	peroxisomes mitochondrium chloroplast
Hodick (1993)	C. globularis (=C. fragilis) protonema from nodes	microscopy	-	cell division nuclei
Sievers and Schröter (1971), Braun and Sievers (1993), Buchen et al. (1993), Braun (1996a), Braun (1996b), Braun and Wasteneys (1998)	C. globularis (=C. fragilis), N. pseudoflabellata protonema, rhizoids	immune-fluorescence labelling sectioning (2-3 µm) slow-rotating-centrifuge-microscope	fluorescein isothiocyanate (FITC)-conjugated anti-mouse IgM fluorescein isothiocyanate (FITC)-conjugated sheep anti-mouse IgG rhodamine-phalloidin in sodium-potassium-phosphate-buffer 2% uranyl acetate and 2% lead citrate acid fuchsin ethanol	stratoliths nucleus microfibrils actin-cytoskeleton myosin-related proteins

Table 2.

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Methods for oospore dormancy breakage for charophyte species from permanent (p) and temporary (t) habitats from different zonobioms after Pott (2005). These are indicated with Roman numbers as given by the literature, superscript labels indicate the zonobiome and are listed below.

Species	Habitat	Zonobiome	Most effective method to break dormancy (storage conditions)	Reference
Chara aculeolata	p	VIII1	+4°C for about 2 months	Forsberg (1965c)
C. aspera	p	VIII	+4°C for about 2 months	Forsberg (1965c)
C. australis	p p	V2 V	wet under semi-natural conditions	Casanova and Brock (1996) de Winton et al. (2004)
C. braunii	p	I3/V	wet storage 22°C	Holzhausen et al. (2022)
C. canescens	p/t p p/t	VI4 VI VI	dry oospores, low temperatures 12 weeks at 10°C in the dark dessication at 5°C	Ernst (1921) Steinhardt and Selig (2011) Holzhausen et al. (2017)
C. contraria	p/t p p	N/A III5,* VII6	drying of oospores wet oospores at cold temperatures (3°C) fresh oospores from mother plant	Shen (1966) Proctor (1967) Sabbatini et al. (1987)
C. corallina	p	V	15°C and darkness for up to 4 days	de Winton et al. (2000)
C. globularis	p p	III* V	dry oospores at cold temperatures (3°C) 15°C and darkness for up to 4 days	Proctor (1967) de Winton et al. (2000)
C. hispida	t	VI	drying of oospores, cold treatments of 1-3 months at 4°C	Holzhausen, unpublished data
	p	VIII	+4°C for about 2 months	Forsberg (1965c)
C. muelleri	p/t	V	dry storage in either dark or ambient light conditions	Casanova and Brock (1996)
C. papillosa	p	VI	dessication at 20°C	Holzhausen et al. (2017)
C. rusbyana	p	III*	dry oospores at cold temperatures (3°C)	Proctor (1967)
C. vulgaris	p p p p t	VIII VIII VIII VIII VI	36 days at 18-22°C wet storage 60 days at 4°C and red light dark at 4°C drying of oospores, cold treatments of 1-3 months at 4°C	Kalin and Smith (2007) Sederias (2003) Sederias and Colman (2007) Sederias and Colman (2009) Holzhausen, unpublished data
C. zeylanica	p p/t p	VIII N/A III*	4°C for about 2 months cold temperatures (5-7°C) for 10 days wet oospores at cold temperatures (3°C)	Forsberg (1965c) Shen (1966) Proctor (1967)
Lychnothamnus barbatus	p	VI	dessication at 5°C	Holzhausen et al. (2017)
Nitella cristata var. ambigua	p/t	V	cold treatment (4-5°C)	Casanova and Brock (1996)
N. furcata	p	VI	4°C and darkness	Sokol and Stross (1986)
	p	V	15°C and darkness for up to 4 days	de Winton et al. (2000)
N. flexilis	p	VIII	247 days sediment storage at 18-22°C, decline of redox conditions in medium	Kalin and Smith (2007)
N. sonderi/N. subtilissima	p/t	V	drying of oospores	Casanova and Brock (1996)

• * sample sites are not given for all strains of the collection
• ¹ coniferous forest land and taiga
• ² warm temperate area with indeciduous forests (laurel forests)
• ³ zone of humid tropical rain forests
• ⁴ temperate, winter cold areas with deciduous forests (= nemoral zonobiom)
• ⁵ zone of the subtropical hot arid semi-deserts and deserts
• ⁶ steppes, semi-deserts and deserts

In 1871, Anton de Bary described the development of the female gametangium up to fertilisation by male spermatozoids (De Bary 1871). Using a Brückean magnifying glass, he was able to observe in detail not only the cell development of the surrounding envelope cells, but also the formation of a tube-like channel, which results from a difference between the convergence of spiral cell tips and cells. Masses of spermatozoids rest near the area of the five tube-like channels, fixed by the gelatinous intercellular compound of the five narrowed channels, allowing the intrusion of spermatozoids as a dense bundle formation to move to the intercellular part above the oocyte. In contrast, the intercellular part is only traversed by a narrowed channel, which ends in a widened funnel shape at the apex of the oocyte. A loose membrane structure allows individual spermatozoids to pass through the apex. Fertilisation is followed by the formation of a thickened cellulose layer and a thickening and change in the colour of the spiral cells. It is thought that the absence of this layer is associated with infertility, although simultaneously the same amount of amylum is present.

The molecular signalling pathways involved during fertilisation or meiosis are still unknown. The exact time of meiosis is still unknown. For a summary, see Guerlesquin and Noor (1982), Schubert et al. (2016), Schubert et al. (In press). Five different opposing hypotheses have been proposed for the exact time point of meiosis in charophytes: (I) just before the germination of the oospore (Oelkers 1916), (II) in the early cell stages of the antheridia and oogonia (Tuttle 1924, Tuttle 1926), (III) during the development of the protonema filament (Gonçalves da Cunha 1936, Gonçalves da Cunha 1942) and (IV) immediately after the fertilisation (Shen 1967b, Shen 1967c). Although hypotheses (I) and (IV) are almost identical, both have been listed because of gaps in the descriptions of cell movement and the resting phase before germination. In 1899, Goertz described the observation of a possible delayed fertilisation by the protrusion of starch into the neck of the female gametophyte (V, Goetz 1899). Despite a comparison with Vaucheria, no further evidence is found in literature (Oltmanns 1895). The myth of meiosis is not well known, mainly due to complications in microtome sectioning. The high starch content of oogonia and oospores makes the procedure difficult. The establishment of methods for ultrathin sectioning of all developmental stages (from oogonia formation to the released oospore) is one of the most important prerequisites for solving this mysterious process.

Cultivation and field studies have shown that the process of fertilisation may be dependent on environmental conditions, such as temperature, light or salinity. Cultivation or germination conditions, as well as species-specific growth behaviour, should be considered when comparing existing literature. For example, Karling (1924) compared the maturation of oospores under different light conditions. Whereas, in greenhouses (high temperature during the day, low temperature at night), the oospores matured, in climatic chambers (high temperature during the day and night), the oogonia remained in an unfertilised state. The same has been proposed for Chara braunii (Sato et al. 2014, Holzhausen et al. 2022). The same is true for Chara aspera with respect to light and salinity. The highest rate of gametangiogenesis was found under natural ecological conditions. Greater variation in conditions compared to the natural ecological status resulted in stress responses such as reduced gametangia numbers or cell death (Blindow et al. 2003). In this context, tracer studies on the polarity and concentration of photoassimilates in the vacuolar sap of different regions of the Chara thalli (sugar-phosphates, carbohydrates (sucrose), malate, amino acids) of C. vulgaris, C. hispida and C. corallina showed an assumed correlation between the accumulation of sucrose and reproduction. For a summary, see Schulte et al. (1994). Briefly, they found out that:

1. differences in sap composition are caused by seasonal changes and reproduction (MacRobbie 1962, Barr 1965, Winter et al. 1987),
2. lateral branches as the source of gametangia have the highest photosynthetic activity (Andrews et al. 1984b, Ding et al. 1991a),
3. gametangia are sinks of photosynthates during reproductive periods (Schulte et al. 1994),
4. a higher activity of 1,5-bisphosphate carboxylase (RuBPCase) and a fourfold higher ferricyanide-dependent oxygen evolution in branchlets than in internodes (Ding et al. 1991b),
5. differences between growing and overwintering periods (Shepherd and Goodwin 1992),
6. intercellular transport occurs via plasmodesmata (e.g. Leitch et al. (1990)) and
7. the intercellular transport is determined by the rate and concentration of cytoplasmic streaming and a possibly unclear “intrinsic regulatory process” (Ding et al. 1991a).

Assuming that the rate of cytoplasmic streaming depends not only on general seasonal changes, but also on the exact environmental conditions, the differences in oospore sizes, as reported by different authors, could probably be explained by the number of cell-cell-connections between branchlets and oogonia and the associated rate of sucrose uptake and starch storage, respectively (Holzhausen 2016). Comprehensive analyses, combining phenological and cytological studies, molecular mechanisms and environmental data over several years including the open question of chloride uptake by oogonia, could resolve this open question.

Asexual reproduction of charophytes

Asexual reproduction by vegetative parts is common in aquatic plants and algae. For reviews about this, see for example Lovett Doust and Lovett Doust (1988), Barrat-Segretain (1996), Barrat-Segretain et al. (1998), Barrat-Segretain and Bornette (2000), Cecere et al. (2011), Agrawal (2012). For most Characeae, vegetative reproduction is a widespread feature, except of species with meteoric appearance. However, there is one exception, Chara canescens, which was never seen to reproduce vegetatively by formation of modified node or rhizoid cells and is, therefore, dependent on the formation of oospores for reproduction (Krause 1997). In addition to its unique position, this species can reproduce parthenogenetically or sexually through dioecious populations and it has been shown that females of parthenogenetic and sexually reproducing populations are genetically distinct (Schaible et al. 2009a, Schaible et al. 2009b, Nowak et al. 2019) and differ in chromosome numbers (Ernst 1917).

In contrast to generative reproduction, there is a limited amount of literature on asexual reproduction. However, only a few scientists (Braun 1852, Giesenhagen 1897, Migula 1898, Bharatan 1987, Sederias 2003, Skurzyński and Bociąg 2011, Pokrzywinski et al. 2020) have studied this type of propagation in detail for species of the genera Nitellopsis, Chara, Lamprothamnium and Paleonitella. According to the time, cell division in nodal cells and the emergence of new shoots were mainly visualised by schematic representations, as studies and visualisation with modern microscopy techniques does not exist, except for the rhizoids themselves (e.g. Braun et al. (1996), Braun et al. (1999)). In 1897, Giesenhagen summarised the differences between bulbils and modified rhizoid nodules, based mainly on the work of Migula (Migula 1898) and his own results (Fig. 2A/B). However, Giesenhagen’s findings are not entirely consistent with Migula’s, as C. aspera rhizoidal bulbils are not unicellular and Giesenhagen gives a detailed description of their formation and cell division (Giesenhagen 1897). The different provisioning of individual rhizoidal cells, reflected in the different sizes within the complex, has not been fully elucidated. The number of modified rhizoid cells is species-specific, with more than four being described as unusual in C. aspera, whereas complexes of three to five or more than 12 have been described for Lamprothamnium macropogon (Braun and Nordstedt 1882, referred as Lychnothamnus macropogon). Fig. 2C shows rhizoidal bulbils of C. aspera with associated microorganisms. The nearest rhizoidal node or the apical part of the cell is assumed to be responsible for the development of new shoots. In his study, Bharathan describes the same type of 'bulbils' for Chara hornemannii, C. aspera, L. papulosum and Lamprothamnium succinctum, but in contrast to earlier studies, they are also referred to as bulbils and not as rhizoid nodules (Bharatan 1987). Interestingly, the author's own results have shown that sterilisation and severance of the connecting rhizoid filaments do not lead to the development of new shoots. So far, no signalling pathways are known for these connecting rhizoid filaments. The study by Bonnot and co-workers showed that the Chara braunii ROOT HAIR DEFECTIVE SIX-LIKE genes do not complement the rhizoid or root-hair development in mosses and land plants, initiating a completely different function (Bonnot et al. 2019). In contrast to this, the induction of germination by the rhizoid/shoot nodule complex is simple and occurs within a few days under laboratory conditions (Giesenhagen 1897, Bharatan 1987, Blindow et al. 2009). Vegetative reproduction can occur via modified shoot nodal cells (nodal bulbils) in addition to modified rhizoid nodes. The ability of these cells to produce new shoots after periods of drought, fragmentation or death of apical cells is well documented (Fig. 2D). This ensures the growth of new populations under unfavourable conditions for oospores (Casanova 1994, Casanova 1999, Blindow et al. 2009). Regeneration of fragmented nodal cells requires the formation of lateral branches or cone-like outgrowths (Giesenhagen 1897) as described for the strawberry-shaped nodal bulbils of Chara baltica, a limited number of new thalli can be developed by a modified node. These are the same as lateral branches, but greatly reduced. Various studies have been published on the structuring of shoot node cells (Braun and Nordstedt 1882, Giesenhagen 1897, Schubert et al. 2016), including cell division (Fig. 2E-G). Interestingly, different forms of overwintering structures can be observed: star-shaped modified shoot nodes in Nitellopsis obtusa (Fig. 2A/B), strawberry-shaped modified nodal cells in C. baltica or cardioid-shaped modified rhizoid cells in Chara fragifera. The existence of star-shaped bulbils in Chara connivens, as mentioned by Clavaud (Clavaud 1863, Giesenhagen 1897), is not fully confirmed. The phenomenon of reduced, starch-filled bulbils acting as reserves can also be found in other species, such as C. baueri, C. vulgaris (= C. foetida), C. globularis (= C. fragilis), C. hispida or C. subspinosa (= C. rudis). Bulbils have also been described in Nitella species from the Southern Hemisphere (De Maisonneuve 1859, Casanova 1994, Casanova 1999). However, the formation and gene expression in these vegetative cells is not fully understood, although a correlation with distribution patterns caused by landmass separation and environmental changes has been hypothesised (Croy 1979). This is supported by barcoding studies that found adaptive mutations of chloroplast and mitochondrial genes in different habitats (Kato et al. 2008, Schneider et al. 2015). The lack of multidimensional approaches, including field, laboratory, microscopic and molecular studies, to address open questions about vegetative reproduction should be addressed in future studies.

Figure 2.

Reproductive or growth units and rhizoid cells of charophytes. (A) fully developed star-shaped bulbil of Nitellopsis obtusa, scale 1 mm. (B) Cell at the beginning of bulbil development, scale 200 µm. (C) rhizoidal bulbil with associated microorganisms of Chara aspera, scale 2 mm. (D) growth unit of Chara filiformis (stem cell), scale 500 µm. (E) characteristic S-shaped division plane of rhizoid cells (*) of Chara vulgaris with additional cells for secondary rhizoids (arrows), scale 1 mm. (F) rhizoid cell of Chara braunii S276, stained with DAPI, scale 200 µm. (G) rhizoid tip of C. braunii S276, stained with DAPI, 200 µm. (H) oospore of C. braunii S276, stained with PI, scale 200 µm (photographs: A. Holzhausen).

Dormancy and germination induction of charophytes

In both permanent and temporary waterbodies, the time between fertilisation and germination is extended by a period of dormancy to increase the possibility of a period of low competition and successful population establishment, although subsequent reproductive capacity is not guaranteed (Casanova and Brock 1990, Vleeshouwers et al. 1995). The adaptive seed trait of temporal loss of germination capacity represents a period of physiological processes, such as mobilising resources and activating/deactivating genes leading to morphological changes and subsequently germination. According to Baskin and Baskin (1985), who described it for land plant seeds, the physiological, morphological and physical state at the time of maturation and germination must be considered and combined with environmental habitat conditions at the time of seed/oospore state changes and during the maturation and germination process.

However, many classification systems have been established because there is no single definition of dormancy (Bewley and Black 1982). The distinction between developmental time, i.e. primary and secondary dormancy, is the simplest and most convenient. While primary dormancy is induced during development and maturation (e.g. Crocker (1916), Hilhorst (1995), Hilhorst (1998), Finch-Savage and Leubner-Metzger (2006)), secondary dormancy is induced only after release from the mother plant and requires the loss of primary dormancy. Interestingly, although not yet confirmed by molecular studies, the hypothetical physiological dormancy model for terrestrial seeds (Hilhorst 1998) also seems to apply to charophytes (Fig. 3). This combined model describes the regulation of secondary dormancy by temperature and the subsequent triggering of germination by the availability of nitrate, light and gibberellic acids. Figure 3 shows the modified model of Hilhorst (1998) and implies that dormancy and germination are directly correlated with the amount of phytochrome receptors present. These red (R) and far-red (FR) receptors act as light sensors in several plants/algae (Inoue et al. 2019), which are mainly located in the nucleolus. Several germination studies have identified that germination is triggered by the supply of gibberellic acid (Sederias and Colman 2007, Holzhausen et al. 2022), although evolutionary studies have shown that the gibberellic acid pathway has evolved in mosses (Nishiyama et al. 2018). Further studies are needed to determine whether chemical precursors could explain this phenomenon. In addition, endogenous gibberellic acid has been measured in studies on C. tomentosa and C. vulgaris (Kaźmierczak and Rosiak 2000, Kaźmierczak 2001, Chowdary 2014). Fertilisation was not found to be related to endogenous gibberellic acid content, but to sex. While the concentration in sterile shoots increases towards the apex, the concentration in fertile shoots declines from the apex downwards. In contrast, the levels in male plants exceed those in female plants.

Figure 3.

Modified hypothetical physiological dormancy model of Hilhorst (1998). Originally this model was developed for seeds of magnoliophytes in soil seed banks, but could be particularly applied on charophytes. I. the stage of maximum dormancy is characterised by lacking phytochrome receptors at the plasma membrane. II. The formation of phytochrome receptors (pr) is temperature-dependent. III. Optimisation of membrane fluidity by increased temperatures. IV. activation of phytochrome receptors by nitrate. V. Formation of photoreceptor complexes by binding of active phytochrome (P_fr). VI. signal transduction for gibberellin synthesis. VII. gibberellin receptor synthesis VIII. gibberellin receptor complex by binding of gibberellin or gibberellin precursors. IX. germination initiation.

The induction of germination itself is a signal for the end of the resting phase (dormancy) and for the presence of favourable environmental conditions for growth. Sensors are used to provide signals on temperature, light, nutrient availability and hydration status. Competitive conditions, on the other hand, are undetectable and represent a 'trial and error' approach. Individual factors, although not always fully distinguishable, are considered in the following subsection.

Growth and in-vitro cultivation

Biomolecular studies on charophytes

Conclusions

In the previous sections, the wealth of studies on reproduction, germination including oospore dormancy and cultivation that can be used on an ad hoc basis and as fundamental resource has been presented. Both the low rates of germination and the difficulty of controlling epiphytic overgrowth have been described in the previous sections. Furthermore, the induction of gametangiogenesis in monoecious and diplostichous Chara species, such as C. hispida, is not successful under artificial in-vitro conditions, as shown by cultures of different populations in different common media (e.g. Forsberg (1965b), Andrews et al. (1984a), Wüstenberg et al. (2011)) and under continuous light cycles of 16 h light to 8 h dark or 12 h:12 h (Fig. 4).

Figure 4.

Cultivation of Chara hispida. While no gametangia could be developed under conditions of common temperature or light regimes and nutritional media, the use of Eco-in-vitro cultivation including the use of logged environmental data from outdoor mesocosm experiments (Holzhausen 2019) is successfully proven for in-vitro gametangia initiation/maturation (photographs: A. Holzhausen).

In contrast, cultivation of C. hispida in outdoor mesocosms at the Rostock University showed that cultivation under natural conditions including temperature, light and continuous oxygenation induced germination, gametangia initiation and maturation (Holzhausen 2019). Logger data of temperature and light intensities including the light-dark cycle were used to identify temperature and light ranges in which the processes of dormancy breakage and germination initiation, gametangia development and maturation take place. Although the decisive factor could not be identified, the transfer of the recorded logger data (temperature, light-dark cycle, intensity, Holzhausen (2019)) to in-vitro cultures showed that the induction of gametangia and maturation of oospores is possible (Fig. 4) using nutritional medium (Wüstenberg et al. 2011).

This Eco-in-vitro-culture method could be particularly useful for species with low in vitro gametangiogenic potential such as C. hispida. In contrast to environmental alga material, these cultures have a drastically reduced microbiome which facilitates the identification of contaminant-specific and bacterial-specific gene sequences. The further understanding and integration of evolutionary, systematic and ecological knowledge will enable the establishent of various in-vitro (and axenic) cultures (axenic strain C. braunii S276, Holzhausen, unpublished) that will allow the identification of reproduction-related gene networks at different reproduction states and of different reproduction modes.

Acknowledgements

Conflicts of interest

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