Phytohormones and Pheromones in the Phycology Literature: Benchmarking of Data-Set and Developing Critical Tools of Biotechnological Implications for Commercial Aquaculture Industry

Rathod, Sachin G.; Bhushan, Satej; Mantri, Vaibhav A.

doi:10.3390/phycology4010001

Open AccessReview

Phytohormones and Pheromones in the Phycology Literature: Benchmarking of Data-Set and Developing Critical Tools of Biotechnological Implications for Commercial Aquaculture Industry

by

Sachin G. Rathod

^1,2

,

Satej Bhushan

¹

and

Vaibhav A. Mantri

^1,2,*

¹

Applied Phycology and Biotechnology Division, CSIR-Central Salt and Marine Chemicals Research Institute, Gijubhai Badheka Marg, Bhavnagar- 364002, India

²

Academy of Scientific and Innovative Research (AcSIR), Ghaziabad- 201002, India

^*

Author to whom correspondence should be addressed.

Phycology 2024, 4(1), 1-36; https://0-doi-org.brum.beds.ac.uk/10.3390/phycology4010001

Submission received: 30 October 2023 / Revised: 16 December 2023 / Accepted: 18 December 2023 / Published: 21 December 2023

Download

Browse Figures

Versions Notes

Abstract

:

Plant hormones and pheromones are natural compounds involved in the growth, development, and reproductive processes. There is a plethora of studies on hormones and pheromones in terrestrial plants, but such investigations are few in the phycological literature. There are striking similarities between the chemical diversity, biosynthetic processes, roles, and actions of hormones and pheromones in both higher angiospermic plants and algae. However, there are substantial knowledge gaps in understanding the genes responsible for hormone biosynthesis and regulation in algae. Efforts have focused on identifying the genes and proteins involved in these processes, shedding light on lateral gene transfer and evolutionary outcomes. This comprehensive review contributes to benchmarking data and essential biotechnological tools, particularly for the aquaculture industry where seaweed is economically crucial. Advanced techniques in plant hormones and pheromones can revolutionize commercial aquaculture by using synthetic analogs to enhance growth, yield, and reproductive control, thereby addressing seasonal limitations and enabling sustainable seedling production. To the best of our knowledge, this is the first comprehensive review that focuses on biosynthetic pathways and modes of action (of five plant hormones and five pheromones), roles (of 11 hormones and 29 pheromones), and extraction protocols (of four hormones and six pheromones) reported in the phycological domain.

Keywords:

aquaculture; biosynthetic pathways; hormones; pheromones; phycology; seedling production

1. Introduction

Algae are a diverse group of photosynthetic organisms that are unrelated ecologically as well morphologically to other groups [1]. They range from microscopic unicellular forms to large multicellular seaweeds, representing a crucial component of aquatic ecosystems, contributing significantly to oxygen production and serving as fundamental sources of nutrition [2,3]. Classified into various taxonomic groups, including green, brown, and red algae, they showcase a remarkable adaptability to diverse environments, highlighting their significance in both freshwater and marine ecosystems [4]. The forms around which considerable trade and economics are developed are marine macroalgae. They represent heterogeneous artificial groups, forms of marine polyphyletic origin, and different evolutionary lineages [5].

The perennial, multilayered seaweed stands of large ‘kelps’ represent the most productive ecosystems, which can sequester significant blue carbon and consequently increase oxygen in the oceanic environment [6]. Seaweeds have a long-standing history of exploitation by humankind for food, fodder, and agriculture, especially in Asia, Polynesia, and South America [7]. Propelled by emerging applications in day-to-day commodity products and biotechnological and medical use, seaweeds have been industrially farmed. In 2018, the commercial production of seaweeds globally reached more than 30 million tons, 97% of which was harvested through aquaculture [8]. Currently, 47 species and two varieties of 27 genera are commercially cultivated—largely in Asian countries [9]. The industry is expected to improve its performance by 12% annual growth and is anticipated to reach USD 30.2 billion by 2025 [10]. The burgeoning global population necessitated substantial improvement in food production even at the cost of limited availability of agricultural land, fast-deteriorating soil quality, shortages of water for irrigation, and protuberances by climate change. It should be noted that, considering current consumption trends, there is an urgent need to produce 50–70% additional food by 2050 [11]. The seaweeds or their extracts have been used since ancient times as soil conditioners or fertilizers in agriculture [12]. Their large-scale application has considerable potential to improve food production [13]. This is because they are needed only in small dosages—often diluted in volume by a factor of 20–500 [14]—as growth stimulants to enhance yield [15], impart disease resistance [16], elevate drought tolerance [17], reduce pest infestation [18], and improve shelf-life of produce [19]. These applications need in-depth scientific understating to unravel their mode of action, which is largely unknown. The empirical evidence showed that the growth responses elicited by seaweed extracts cannot be attributed to the presence of macro- and micro-elements alone, but plant growth regulatory compounds might also play a catalytic role [20]. The discovery of different classes of hormones in seaweeds or their extracts was evident from the work that was carried out during the late 1960s to early 1970s [21].

This comprehensive review deals with the critical aspects of phytohormones and pheromones reported in algae (encompassing both macro- and microscopic forms). It is well evident that plant hormones play a pivotal role in regulating growth and development while pheromones are needed for induction of reproduction and sexual maturity in algae. Further, we tried to elucidate on how their biosynthetic pathways are similar to those in higher angiospermic plants, e.g., response to climate change and stress from the herbivores. We have tried to figure out the detailed ways these substances work in macroalgae as well as microalgae, especially in their defense mechanisms. If we understand this process better, it could help us to have effective control over algal growth and dealing with infestations. The study also highlights the importance of understanding the reproduction of commercially valuable macroalgae and microalgae, suggesting that hormones and pheromones may play a crucial role in advancing spore-based cultivation technologies. We believe that the data synthesis provided here would be useful in developing critical tools of biotechnological implications for the commercial aquaculture industry.

2. Hormones and Pheromones Reported in the Phycology Literature

Phytohormones are produced within the plant cells at extremely low concentrations. They act as signaling molecules and control almost all aspects of development and growth. The hormones acquire a greater significance as messenger or signaling molecules, where contact between adjacent cells plays a key role in regulating metabolism at the tissue level [22]. Eleven different plant hormones have been discovered in algae (Table 1), i.e., auxin, cytokinin, gibberellin, abscisic acid, ethylene, rhodomorphin, jasmonic acid, brassinosteroids, salicylic acid, strigolactones, and polyamines. More recently, strigolactones have been identified from freshwater alga Chara coralina as a new branching inhibiting hormone [23]. Polyamines, e.g., thermospermine, are considered to be one of the plant hormones and are present in various seaweeds like Ascophyllum nodosum, Fucus vesiculosus, and Sargassum horneri (Table 1).

The concept and different terminologies related to hormone research in the phycology literature have been adopted largely from higher plants. The studies on hormones in algae are uncoordinated and not as advanced as in higher plants. This review primarily utilizes the literature on seaweeds and other algal forms like microalgae (of marine as well as freshwater origin) wherever necessary to substantiate the phycological origin of data. The presence of gibberellin activity was first reported in Enteromorpha prolifera (now Ulva prolifera) and Ecklonia radiata [55]; auxin activity in Ulva pertusa (now Ulva australis), Undaria pinnatifida, and Hizikia fusiformis (now Sargassum fusiforme) [56]; and cytokinin activity in species of Laminaria and Fucus [57]. Even the concentration of hormones like IAA has been studied in zygotes and mature tissues of Fucus distichus, which was 2–9 ng g⁻¹ fr wt and is in a slightly lower proportion than in higher plants [58].

The word pheromone is derived from a Greek word that means ‘to carry’, which signifies its role as it carries information regarding the availability and favorability of conditions for the organism to sexually reproduce [59]. There have been a number of studies related to the bioactive metabolites and hormones produced by seaweeds that can effectuate interspecific signaling, but very little is known about the chemical cues that affect the members of the same species, i.e., interspecific interaction of the seaweeds. These cues are the pheromones, which can help in deciphering the factors that induce sexual reproduction in those seaweeds. A number of pheromones have been identified in various seaweeds (Table 2).

3. Biosynthetic Pathways

3.1. Hormones

3.1.1. Auxin

Auxin (IAA) biosynthesis occurs mainly by two pathways, i.e., tryptophan-dependent (Figure 1) and tryptophan-independent (Supplementary Figure S1). The tryptophan-dependent pathway follows four different routes: indole-3-pyruvic acid (IPA) pathway, indole-3-acetaldoxime pathway, tryptamine pathway, and IAM pathway, of which indole-3-pyruvic acid (IPA) pathway and tryptamine pathway (TAM) are the main routes for IAA biosynthesis in the plant [77]. In algae, the tryptamine pathway (TAM) could be the most probable pathway for auxin biosynthesis. The tryptophan decarboxylase enzyme has been reported from microalgae Chlamydomonas reinhardtii [78]. Algal counterparts of many auxin biosynthetic enzymes from higher plants like C-S lyase, and nitrilases have also been reported in Ectocarpus siliculosus, Ostreococcus lucimarinus, Micromonas pusilla, Chlorella variabilis, Volvox carteri, etc. [78]. Amino acid sequence comparison of the Flavin-containing mono-oxidases, YUCCA and FLOOZY, involved in auxin biosynthesis has been carried out between algae like Ectocarpus siliculosus, Ostreococcus lucimarinus, Ostreococcus tauri, and Chlorella variabilis and higher plant Arabidopsis. Their comparison has revealed homology with a high confidence value, suggesting parallels in the biosynthetic pathways of both groups [78]. Similar analysis performed for the enzyme tryptophan aminotransferase did not result in any sequence similarity, suggesting the absence of the indole-3-pyruvate pathway of auxin biosynthesis in algae [78].

3.1.2. Cytokinins

In higher plants, isoprenoid cytokinin biosynthesis takes place via two different pathways. The direct route includes the formation of N6-isopentenyladenosine monophosphate (iPMP) from AMP and pyrophosphate, catalyzed by isopentenyltransferase (IPT) (Figure 2) [80]. In the second pathway, the synthesis of isoprenoid cytokinin takes place by making changes in the structure of tRNA containing cis-zeatin [30]. This pathway can be found in all organisms except Archea [81]. In marine macroalgae (chlorophyta, pheophyta, and rhodophya), both isoprenoid and aromatic cytokinins and their conjugates have been detected, indicating the presence of complex inter-conversion systems and regulation of their activities, but this second indirect pathway, which includes tRNA degradation, seems to be a characteristic feature of marine macroalgae [30]. Cytokinin biosynthesis has been reported from different macro- as well as microalgae. The macroalgae include Cladophora capensis, Ulva fasciata, Caulerpa filiformis, Sargassum heterophyllum, Porphyra capensis, Amphiroa bowerbankii, Dictyota humifusa, etc. [30,36], and the microalgae include Protococcus viridis, Chlorella minutissima, Chlorella sp., and Scenedesmus sp. [82]. Interestingly, isopentenyladenine (iP) and cis-zeatin (cZ) forms have been detected in higher concentrations than dihydrozeatin (DHZ) conjugates; whereas, no N-glucosides have been reported from various marine macroalgae like Ulva, Caulerpa, Sargassum, Macrocystis, Porphyra, Hypnea, Amphiroa, etc. This suggests that they may have the tRNA-dependent pathway as the preferred route for cytokinin biosynthesis [83]. On BlastP search of amino acid sequence of biosynthetic enzymes like isopentenyltransferase from higher plants, a strong homology with proteins from various algal species like Ectocarpus siliculosus, Volvox carteri f. nagariensis, Micromonas pusilla, and Chlorella variabilis was observed. It could be concluded that the cytokinin biosynthesis in microalgae (Micromonas, Chlorella, Ostreococcus, Volvox, Thalassiosira, and Phaeodactylum) and macroalgae (Ectocarpus siliculosus) possibly occurs via pathways similar to higher plants [78].

3.1.3. Gibberellins

GAs are chemically diterpenes, and their synthesis, at the early stages, takes place via one of two pathways of isoprenoid biosynthesis, i.e., through the mevalonic acid pathway or methylerythritol phosphate pathway [85]. In higher plants, GA is mostly synthesized through the methylerythritol phosphate pathway, which occurs in plastids [86]. The main reaction of GA biosynthesis is the cyclization of geranylgeranyl pyrophosphate (GGPP) into copalyl pyrophosphate and the final conversion into ent-kaurene (Figure 3). The reactions are catalyzed via the copalyl pyrophosphate synthase (CPS) and ent-kaurene synthase, respectively. Protein sequences similar to the Arabidopsis enzymes, copalyl pyrophosphate synthase, ent-kaurene synthase, and ent-kaurenoic acid oxidase have not been reported from algae [78]. The reason may not be the absence of corresponding enzymes, but the shortage of the available proteomic data. GA-20 oxidase enzyme has been characterized in a green alga Chlamydomonas reinhardtii. The BlastP search of this enzyme showed a considerable homology to A. thaliana sequence of late-stage enzymes of GA synthesis. This suggests that the GA biosynthesis pathway in algae may not differ too much from the higher plants. At the same time, it is also clear that more research-based evidence is required to understand the exact pathway in algae [78].

3.1.4. Abscisic Acid (ABA)

ABA biosynthesis occurs through the precursor isopentenyl pyrophosphate or directly via the degradation of carotenoids (Figure 4). Carotenoid synthesis is the first stage of ABA biosynthesis where all the isoprenoids and carotenoids are produced from isopentenyl pyrophosphate (IPP) [87]. For the carotenoid synthesis, geranylgeranyl pyrophosphate (GGPP) is formed from the isopentenyl diphosphate (IPP), and GGPP is converted into phytoene, which is catalyzed by phytoene synthase (PSY). Phytoene desaturase (PDS) catalyzes the conversion of phytoene into ζ-carotene and sequentially converts it into lycopene, β-carotene until the zeaxanthin [78], or via the direct pathway in which zeaxanthin forms from the isopentenyl pyrophosphate (IPP) via farnesyl-diphosphate [84,88]. The first key reaction of ABA biosynthesis is a conversion of zeaxanthin into trans-violaxanthin via two-step de-epoxidation catalyzed by zeaxanthin epoxidase (ZEP). The trans-violaxanthin is then converted into 9-cis-neoxanthin by the enzyme neoxanthin synthase. After that, xanthoxin is synthesized by the oxidative splitting of 9-cis-violaxanthin and/or 9-cis-neoxanthin, and the reaction is catalyzed by the 9-cis-epoxycarotenoid dioxygenase (NCED) [78,84]. Finally, there are three possible ways for the last step of ABA biosynthesis from xanthoxin, i.e., three different intermediate compounds can be formed before finally yielding ABA. These intermediates are ABA aldehyde (as shown), xanthoxinic acid, or abscisic alcohol [89].

ABA biosynthesis in the unicellular green alga C. reinhardtii occurs via a neoxanthin-mediated pathway [90]. It has been reported that 9′-cis-neoxanthin, a suitable substrate for ABA production, was present in the green algae containing chlorophyll a and b, whereas 9′-cis-neoxanthin was not found in other algal divisions, such as Heterokontophyta and Rhodophyta. However, all of these organisms do synthesize ABA, which could very well be via the direct pathway involving farnesyl-diphosphate [88]. Furthermore, the inhibition of known carotenoid precursors did not affect the ABA accumulation in cyanobacteria, Dinophyta, and Rhodophyta, suggesting either a new or unknown carotenoid precursor or a direct route of ABA biosynthesis from IPP precursors, which needs to be studied in detail [87]. Since the preliminary steps of ABA biosynthesis occur with those of carotenoid biosynthesis, it may be anticipated that homogeneous enzymes catalyzing these reactions are present in a wide set of algae. In the BlastP results of amino acid sequences of enzymes PSY, PDS, and ZEP from A. thaliana, a considerable number of similarities were found with diverse algae. Similarly, the homologs of enzymes NCED and xanthoxin dehydrogenase like SDR (dehydrogenase/reductase, which is involved in spontaneous rearrangement from xanthoxin to ABA aldehyde) specific for ABA biosynthesis were also identified in different algae, i.e., Ectocarpus siliculosus, Chlamydomonas reinhardtii, and Chlorella variabilis. It can therefore be concluded that the algal representatives contain homologous enzymes participating in ABA biosynthesis since its biosynthetic pathway in algal representatives is similar to that in higher plants [78].

3.1.5. Ethylene

Ethylene formation in marine algae was first reported by Watanabe and Kondo [91]. Methionine acts as an efficient precursor for the dimethylsulphoniopropionate (DMSP) (dimethyl-β-propiothetin) biosynthesis. DMSP is converted into acrylate and dimethyl sulfide by the unknown enzyme protein (now, DMSP lyase). Acrylate is finally converted into the ethylene by the action of acrylate decarboxylase (as yeast decarboxylase) (Figure 5) [91]. The different steps in this pathway were reported in numerous algae Ulva lactuca, Polysiphonia fastigiata, Pyropia tenera (as Porphyra tenera), Ulva pertusa, Codium fragile, Laminaria sp., Fucus vesiculosus, and Digenea simplex [91]. A similar pathway of ethylene biosynthesis was also reported in Ulva intestinalis, which is initiated with the methionine and converted into dimethylsulphoniopropionate (DMSP) by the enzyme methionine transaminase, and further, DMSP lyase converts DMSP to acrylate and acrylate is then finally converted into ethylene via action of enzyme acrylate decarboxylase [92]. In contrast, a different pathway of ethylene biosynthesis was reported in unicellular green algae (Haematococcus pluvialis), which is similar to higher plants. In H. pluvialis, ethylene biosynthesis initiates with L-methionine as precursors, and further, methionine is converted to S-adenosylmethionine (SAM/AdoMet), 1-aminocyclopropane-1-carboxylic acid (ACC), and finally to ethylene via the action of ACC oxidase. The enzymatic complex of the last step of ethylene biosynthesis to ACC oxidase differs from the higher plants. In Haematococcus pluvialis, this enzyme is stimulated by Co²⁺, Mn²⁺, and Ag²⁺, inhibited by Cu²⁺, salicylhydroxamic acid, and by dark, while not affected by Zn²⁺, Fe²⁺, or Mg²⁺. In plants, this enzyme is stimulated by Fe²⁺, Mn²⁺, or Cu²⁺ and inhibited by Co²⁺ [93]. Intermediate compound ACC treatment increases the ethylene production in the chlorophytes Haematococcus pluvialis and Ulva intestinalis, and the red algae Pterocladiella capillacea, like in the higher plant [90]. Therefore, seed plants, red and green algae convert ACC to ethylene, and this pathway is consistent and conserved throughout the plant kingdom [94].

3.2. Pheromones

Pheromones are sex hormones involved in the highly synchronized and regulated process of induction of reproduction in algae [71]. The paucity in the knowledge of exact life cycle stages and reliable methods to induce sexual reproduction are major impediments in identifying the pheromones. These things considered, direct structural elucidation is cumbersome because of the minuscule quantities at which these compounds are secreted in cultured uni-algal samples [95]. The pheromones produced by algal cells act as chemoattractants, facilitating recognition of motile gametes of the opposite sex, thereby enhancing fertilization efficiently [74]. The diversity of pheromone and signaling systems showed considerable diversity and complexity, both within and between algal groups [96]. This section deals with types, biosynthetic pathways, roles, and modes of action of pheromones reported from seaweeds. The pheromones reported from brown, green, and red algae are described below.

3.2.1. Giffordene

Giffordene (2Z,4Z,6E,8Z)-undeca-2,4,6,8-tetraene) has been isolated from Feldmannia mitchelliae (as Giffordia michellae) gyno-gametophytes. Eicosapentaenoic acid is the precursor of giffordene biosynthesis via hydroperoxide HPEPE as an intermediate. The enzymatic carboxylation of 3Z,6Z,9Z-dodecatrienoic acid forms 1,3Z,5Z,8Z-undecatetraene, which ultimately shifts 1,7-sigmatropic hydrogen to obtain giffordene (Figure 6) [97].

3.2.2. Dictyotene

Ectocarpus siliculosus, secretes dictyotene along with ectocarpene, hormosirene, and finavarrene, which act as chemoattractants for male gametes [99]. When E. siliculosus female gametes were externally supplemented with arachidonic acid, there was de novo synthesis of dictyotene (6-butylcyclohepta-l,4-diene), and undeca-(1,3E,5Z)-triene was observed (Figure 7) [100].

3.2.3. Cystophorene

Cystophorene fits in the class of organic compounds as alkatrienes. These are acyclic hydrocarbons that contain exactly three C:C double bonds [101]. Cystophorene (<1%) was also found to be released in trace amounts from suspensions of female gametes of Ectocarpus siliculosus along with ectocarpene (>95%) and dictyotene (ca. 3–4%). First, the fatty acid is activated to the (9S)-hydroperoxide-(9S) HPETE via lipoxygenase. It is followed by an oxidative breakdown into the polar fragment 9-oxonona-5(Z), (E-)dienoic acid, and a respective hydrocarbon (Figure 7) [102].

Figure 7. Biosynthesis of dictyotene and cystophorene (C11 hydrocarbons) in brown algae Ectocarpous siliculosus [103].

3.2.4. Ectocarpene

Ectocarpene is biosynthesized in numerous species of brown algae and was the first isolated pheromone in Ectocarpus siliculosus [61]. PUFAs, such as 9-hydroperoxyeicosatetraenoic acid (9-HPEPE) in the brown algae, are converted to biosynthesized ectocarpene via the consecutive catalytic action of lipoxygenase and hydroxyperoxide lyase (Figure 8) [102].

3.2.5. Dictyopterene

Dictyopterenes are characteristic volatile substances, which are the main constituents of oceanic odor. Additionally, some gamete-attracting substances and flavors, which is a characteristic feature in brown algae, also consist of dictyopterenes, which are mainly C11 hydrocarbon compounds [106]. Neodictyoprolenol [(3S,5Z,8Z)-1,5,8-undecatrien-3-ol; (3S)-1] was assumed to be a possible reaction intermediate of the biosynthesis of the pheromones in the brown seaweed [61,107]. The stereospecific shifting of the hydroxyl group from (9,S)-hydroperoxides (Carbon-9 to Carbon-12) via a six-membered ring ultimately leads to the formation of dictyopterenes (Figure 9).

4. Role of Hormones in Algae

4.1. Auxin

Auxin, found in higher plants, algae, microorganisms, fungi, and animals [108,109], plays a key role in plant growth and development. Generally, the hormone concentration found in algae is much lower as compared to the higher plants [110]. Its function in the growth and development of algae is similar to that in higher plants [111]. Auxin plays a key role in cell division and elongation, suppresses branching at the apical and intercalary regions in red algae Grateloupia dichotoma [112], and has a significant role in the determination of zygote polarization in fucoid algae, i.e., Fucus distichus and Fucus vesiculosus [58,113]. Supplementing the axenic culture of Ulva lactuca germlings in enriched seawater with kinetin and IAA resulted in the formation of a normal flat blade, which further increased the length of the filament with the addition of gibberellin [26]. Similar results were also found with different algae, like Fucus spiralis, Porphyra tenera, and Enteromorpha compressa, when exogenously supplied auxins, p-hydroxy-phenylacetic acid (OH-PAA), and PAA, inducing branching and broadening of fronds [24,114,115]. Inhibition of apical dominance was also found in macroalgae when the apical meristem was removed or damaged, thereby inducing the growth of axillary buds and the formation of lateral branches [116]. IAA concentration in Caulerpa was in the same range as in angiosperms [117]. The activity of IAA in cultured Caulerpa triggered the initiation of leaf-like structures and slower elongation of rhizome-like structures [28]. The synergistic effect of IAA, kinetin, and gibberellic acid (GA) studied in Ulva lactuca induced significant growth higher than each of these hormones individually [26]. The role of IAA in tissue differentiation in multicellular algae has been evident in the literature, in addition to the role in cell elongation and cell division, as observed in higher plants [29]. IAA application also induces cell division by upregulating the genes that encode CDKs, Cycs, CDCs, and tubulins, resulting in increased branch number and promotion of rhizoid branching in Gracilariopsis lemaneiformis [118].

4.2. Cytokinins

Cytokinins regulate key processes like cell division and growth activation in algae just like in higher plants [82]. Kinetin (cytokinin) shows a positive effect on the growth of thallus when added simultaneously with GA, resulting in the formation of adventitious branches from the apical portion of Fucus vesiculosus [34]. Independently as well, kinetin or GA can partly replace the apical cells in Sphacelaria furcigera and increase the length of newly formed lateral branches (apical dominance) from the injured parts of the alga [31]. Cytokinins show less diversity in algae than higher plants but they perform a vital role in the growth and morphogenesis of algae [85]. Cytokinin-like activity occurred in Sargassum heterophyllum during gamete release and at the beginning of receptacle development [119]. Intercalary meristem of young blades of the Macrocystis pyrifera exhibit cytokinin activity in the form of free bases or ribosides and are responsible for cell division, whereas older blades contained cytokinins in the form of O-glucoside as a storage form [120]. Cytokinin, present in seaweed, in the form of free bases and ribosides are the physiologically active forms and can be detected in low concentrations, as they are actively utilized in several developmental processes [36]. Cytokinins, play an important role in the early growth of the receptacles in Sargassum muticum and are also responsible for thalli to possess mature spermatangia and carpogonia in Porphyra perforata [33].

4.3. Gibberellins (GAs)

The regulatory action of GA is well studied in higher plants; however, very few studies have been undertaken to understand their role in algae. In Fucus spiralis and Tetraselmis sp., just like the higher plants, the gibberellins significantly contribute to inducing tissue differentiation via cell elongation and cell division [29]. Such GA-like activities were also reported in Fucus vesiculosus, F. spiralis (Phaeophyceae) [121], and Caulerpa paspaloides (Chlorophyta) [122]. GA treatments of red and brown algal cultures can induce branching and control the growth of axial structures similar to the higher plants. GA3 increases the number of antheridial filaments and spermatids in Chara vulgaris, while the anti-gibberellin, in this case, AMO-1618, inhibits its effect [35]. Exogenous application of GA3 distinctly increases the number of adventitious branches formed on fragments from the apical parts. GA3 also shows a positive effect, in combination with kinetin, on the growth and regeneration of Fucus vesiculosus [34]. In contrast, GA3 inhibits the morphogenesis in the tissue culture of the red alga Grateloupia doryphora [49].

4.4. Abscisic Acid (ABA)

In higher plants, abscission of buds and leaves and dormancy in seeds is caused by ABA and can also inhibit its growth. ABA can be detected in higher concentrations during stress conditions in vascular plants [123]. Similarly, ABA can also be found in many algal groups during stress conditions [124]. However, the concentrations in algal cells are lower as compared to the higher plants [88]. Exogenous ABA accelerates sorus development in addition to its role as a suppressor of vegetative growth in the brown alga Laminaria [125]. In Laminaria, ABA regulates the transition of sporophyte from growth to the stage of propagation [78]. ABA levels in Dunaliella parva, Draparnaldia mutabilis, and Dunaliella acidophila increase with the increase in salinity and pH of the culture media [38,124]. Therefore, changes in endogenous ABA levels due to different environmental conditions may provide pieces of evidence for their possible roles in algae. Higher ABA in Ulva fasciata was observed when collected from a rock pool of the upper intertidal zone since it was more exposed to adverse conditions as compared to Dictyota humifusa, which was collected from a mid-intertidal zone. Therefore, ABA acts as a stress hormone in seaweeds and performs a role in growth inhibition [36].

4.5. Ethylene

In higher angiospermic plants, ethylene biosynthesis occurs during the ripening process, in which biosynthesis may be activated via IAA or by any other physiological stress [84]. Ethylene promotes cap production in Acetabularia acetabulum (as Acetabularia mediterranea) [41], and its precursor 1-aminocyclopropane-l-carboxylicacid (ACC) promotes cell division and cap development in Neoporphyra perforata (as Porphyra perforata) [25]. During sexual reproduction, ethylene regulates gamete formation and protects against stress-induced damage in Neopyropia yezoensis, whereas its precursor, 1-aminocylopropane-1-carboxylic acid, regulates sexual reproduction by inducing the gametophytes to form spermatangia in Neopyropia yezoensis [126,127]. Ethylene also plays a key role in cell wall metabolism, photosynthesis, and abiotic stress responses in Spirogyra pratensis [128].

4.6. Brassinosteroids (BRs)

Brassinosteroids are polyhydroxylated steroid hormones with ubiquitous distribution that regulate the growth and development of higher angiospermic plants. The first report of Brassinosteroids viz, brassinolide (BR), and castasterone (CS) from algae was from the extract of Ecklonia maxima [42]. BRs stimulate the cell division and growth in Chlorella vulgaris, mostly influencing the number of algal cells, phosphorus, chlorophyll, and monosaccharide content in this alga [129]. In Chlorella vulgaris, BRs can regulate protein and lipid content, thereby enhancing the energy storage capacity of the alga during stress conditions like high temperatures [130,131]. It can also boost the stress-responsive ABA content with temperature rise [132]. BRs also play an important role during stress and defense, either individually or along with the primary defense hormones [133].

4.7. Jasmonic Acid (JA)

Jasmonic acid is one of the primary plant defense hormones in addition to SA and ethylene [134]. JA and its derivatives play a vital role as hormones and can induce defense responses by producing oxylipins (defense mediators) and prostaglandins (defense chemicals against grazers) in the red macroalga Chondrus crispus [44]. However, contradictory observations have been made, which suggested that JA- and methyl jasmonate (MeJA)-like compounds may be active just in higher plants and do not play any role in the algal defense system. JA and MeJA may not be ubiquitous in all red algae, as none were detected in the Gracilaria chilensis even after exposure to pathogen attack [135]. Therefore, the role of JA in algae needs to be re-evaluated extensively using the latest analytical techniques on various algal taxa.

4.8. Polyamines (PAs)

The polyamine biosynthetic pathway is conserved in bacteria, animals, and higher angiospermic plants [136]. Polyamines play an important role in physiological metabolism, which eliminates the active oxygen-free radicals, giving the plant tolerance to oxidative stresses [137]. Consequently, polyamines like putrescine and spermidine can help in the acclimation due to the oxidative stress caused by hyposaline conditions in green macroalga Ulva fasciata [47]. Similarly, in the red alga Grateloupia doryphora, during hyposaline shock, the level of putrescine, spermidine, and spermine rises, which has been attributed mainly to the decrease in transglutaminase activity [48]. Exogenous application of PAs can lead to effects similar to 2,4-D in Grateloupia and plays an important role in the development of cystocarp and in the release and development of spores in cultivated species of red macroalga Grateloupia sp. [49,51]. Putrescine and spermidine also play important roles in the transformation of the carposporelings into cell masses that produce shoots. Furthermore, the combination of putrescine, spermidine, and spermine leads to the formation of bigger sizes of cell masses and ultimately to a higher amount of shoot per cell mass [49]. These three are ubiquitous aliphatic amines that are also involved in reproduction in higher angiospermic plants and algae. A higher level of PA (putrescine) in immature cystocarps as compared to mature cystocarps of Crassiphycus corneus (as Gracilaria cornea) was observed, which declined in the transition event of reproduction from the infertile to the fertile state [52,53]. These reports suggest the involvement of polyamines in the reproductive events and other cellular processes in the algae as well.

4.9. Salicylic Acid (SA)

Salicylic acid (SA) is best known for mediating host responses against pathogen infection as it plays an important role in eliciting the defense responses [138]. Some evidence shows that SA plays an important role in the oxidative defense for protection against environmental stresses in seaweeds in a similar way as found in higher plants. SA treated Saccharina japonica (as Laminaria japonica) sporophytes before heat stress improved their thermotolerance by altering antioxidant enzymatic activity with increased superoxidase dismutase (SOD), peroxidase (POD), and catalase (CAT) activity [79].

4.10. Strigolactone (SL)

Strigolactones (SLs) are the newly categorized phytohormones that regulate plant growth, development, and metabolism. Strigolactones have been linked to a variety of physiological processes such as seed germination, nodulation, inhibition of bud outgrowth and shoot branching, photomorphogenesis, and physiological responses to abiotic stimuli [139]. SLs are present in basal Embryophytes, where they are involved in signaling role promoting the arbuscular mycorrhizal (AM) symbiosis [140]. This hypothesis, however, can be challenged by the fact that Charales, which do not participate in AM symbiosis, also synthesize and exude SLs into the medium. Such evidence, supported by sequence and metabolite profile, concluded that the widespread existence of SLs in the green lineage was probably more hormonal than symbiotic [23]. The closest freshwater green algal relatives of land plants, Charales, produce and exude strigolactones, which help them to survive fungal colonization. It was also proven experimentally that the exogenous SLs stimulate rhizoid elongation in Chara coralina [23]. Based on the literature survey, it has been suggested that strigolactones are not found in marine macroalgae. But, the lack of complete genome sequences for lower-order plants, including marine macroalgae, may make such assumptions difficult [141]. However, the strigolactones have been found in the liquid seaweed extract (Seasol™, Seasol International Pty Ltd, Bayswater, Australia), which is made from the biomass of Durvillaea potatorum and Ascophyllum nodosum, which have extensive applications in agriculture [142].

4.11. Rhodomorphin

Rhodomorphin is produced by rhizoidal cells in Griffithsia pacifica, a red alga [139], and is a species-specific growth regulator [21]. A study on Griffithsia sp. revealed its role in the repair of rhizoids, decapitated filament, and its elongation, but no such role has been observed in shoot cell repair [39,143].

5. Role of Pheromones in Algae

5.1. Sporulation Inhibitors

Axenic culture of Ulva mutabilis produces two such inhibitors. Sporulation inhibitor-1a (SI-1a), which is a glycoprotein, is produced by their cell wall, and Sporulation inhibitor (SI-2), which is a non-protein, is produced in the space between the two blade cell layers. Both SI-1 and SI-2 play important roles in keeping the thalli in a vegetative state by suppressing gametogenesis. The absence or removal of these sporulation inhibitors causes induction in gametogenesis from the mature blades in Ulva mutabilis [60].

5.2. Swarming Inhibitors

Swarming inhibitors act as regulatory factors during gametogenesis and are excreted during the determination phase of the gametes and can inhibit the gamete formation event in Ulva compressa (as Ulva mutabilis) [60,144].

5.3. Ectocarpene

Ectocarpene is a chemoattractant hydrocarbon released by female gametes to attract its male counterparts [61]. In most of the brown algae, the fertilization is boosted by such chemical messengers. Ectocarpene is also released by female gametes of Chorda tomentosa to attract male gametes [69]. Ectocarpene is the first reported pheromone in brown algae Ectocarpus siliculosus and is known to induce chemokinesis. It has also been reported in Sphacelaria rigidula, Adenocystis utricularis [63], and in Ectocarpus fasciculatus [62].

5.4. Dictyotene and C11 Sulfur Compounds

Dictyotene and other C11 compounds generally found in brown algae can perform functions like saving the spores, zygotes, and germlings against mesograzers like amphipods [68]. These compounds and their free products also play an essential role in the chemoattraction of gametes in addition to keeping the mesograzers away from the developing zygotes. These are the volatile compounds, reported mainly in Dictyota diemensis, Dictyota dichotoma, Dictyopteris membranacea, D. delicatula, and Sargassum filipendula [64,65,66,67]. Volvox is reported to produce protein erogens; similarly, Allomyces and brown algae have been reported to secrete terpenoids and hydrophobic hydrocarbons, respectively. In Dictyota diemensis, dictyotene has also been reported to act as erotactins, the compounds attracting sperms [74].

5.5. Ochtodene

It is a monoterpene pheromone reported from the Ochtodes secundiramea, which protects this alga against predation by many rapacious herbivores. It also has antibacterial activity against Staphylococcus aureus. Therefore, its antimicrobial role is also important [76].

5.6. Other Chemoattractants

Most of the pheromones perform similar kinds of roles during sexual reproduction. Some of the pheromones in seaweeds and their discovery as chemoattractants are given in Table 2.

6. Mode of Action of Hormones in Algae

Certain plant/algal hormones, unlike animal hormones, have multiple physiological functions [145]. They are produced in cells and then bind to specific receptor proteins to carry out downstream signaling. Their liaison results in a change in cell function and the activation of a signal transduction pathway. The concentration of individual hormones is not important, but the response of hormones is usually governed by the sum effect of other hormones either in tandem or vice versa [146].

In higher angiospermic plants, auxin signal may be perceived at the extracellular matrix, at ER, or inside the nucleus with the help of a receptor ABP1 (Auxin-Binding Protein1) [147,148]. ABP1 homologs have also been found in genomes of Chlorella variabilis NC64A, Chlorella pyrenoidosa, and Chlamydomonas reinhardtii. These proteins form the auxin-binding pocket, which in the presence of auxin, induces transcription of auxin responsive genes [149]. These findings suggest the early emergence of a primitive form of auxin receptors in microalgae [150]. Additionally, more genome sequences of a variety of algae are required to elucidate the origin of auxin signaling in them.

Cytokinin signaling involves the phosphorylation of cytokinin, which binds to the extracellular portion of cytokinin response1 (CRE1), known as the CHASE domain, localized at the plasma membrane [84]. The cytokinin signaling components have evolved in microalgae, and further analogous evolution occurred among different algal lineages [150]. In Arabidopsis, cytokinin is perceived by AHK receptors located in the endoplasmic reticulum, triggering their histidine kinase activity [151]. These receptors are also common in the algal genome [152]. This histidine kinase activity leads to a cascade of phosphorylation from the cytoplasm to the nucleus, ultimately activating the transcription of type-A Arabidopsis Response Regulators (ARRs) and CRFs. Homologous components of these proteins (type-B Arabidopsis Response Regulators and Histidine-Containing Phosphotransmitter 1) have also been found in green microalgae (Nannochloropsis oceanica), suggesting the similarities in their mode of action in microalgae (Nannochloropsis) and plant (Arabidopsis) [152]. The phosphorylated type-A ARRs then interact with various effectors to bring about cytokinin responses [84,151].

Gibberellic acid (GA) molecules bind to the GID1 (GIBBERELLIN INSENSITIVE DWARF1) receptor in higher angiospermic plants, which then interacts with DELLA proteins [153]. DELLA proteins interact with DNA-binding proteins, which are regulated by PIFs (Phytochrome-Interacting Factors) [154]. GID1 receptor orthologs have been identified in microalgae via the functional motif analysis and revealed that the GID homologs have the catalytic triad (S, D, and H) of the hormone-sensitive lipase (HSL) family in microalgae [150]. Therefore, this supports the inheritance of GA signaling from microalgae, which might be the crucial source for the foundations of the higher plant hormone systems. However, some proteins (DELLA and the F-box protein SLEEPY1) involved in mediating GA signaling have been found only in land plants and not in microalgae [155]. This warrants a thorough exploration of downstream signaling molecules involved in GA [150].

Three abscisic acid (ABA) receptors have been identified in higher angiospermic plants, i.e., chloroplast envelope-localized ABA receptor (ChlH/ABAR), plasma membrane-localized GTG1/GTG2 (GPCR-type G protein 1 and 2), and nucleo-cytoplasmic PYR/PYL/RCARs (pyrabactin resistance/pyrabactin resistance-like/regulatory component of ABA receptors) [156,157]. Among these, the nucleo-cytoplasmic PYR/PYL/RCAR receptors are considered the most established ABA perception receptors. These receptors have not been identified in microalgae to date, even though the downstream phosphatases (SNF1-Related Protein Kinase 2) of the ABA signaling pathway are conserved from microalgae to higher plants [158]. Nevertheless, ABA-related genes in algae have not been explored, and it is necessary to compare them to gain more definite proof of the evolutionary origin of ABA-related genes [159].

Ethylene perception in higher angiospermic plants occurs through membrane-bound receptors embedded in the endoplasmic reticulum (ER). Arabidopsis has five known ethylene receptors, which are ETR1 (Ethylene Response1), ETR2, ERS1 (Ethylene Response Sensor1), ERS2, and EIN4 (Ethylene Insensitive 4) [160]. Ethylene receptor complexes, comprising ETR1, ERS, and EIN4, have been widely reported in microalgae (Micromonas sp.) [150]. In addition to that, ethylene binding sites have also been confirmed in a cyanobacterial protein [161].

In silico genome-wide homology search analysis revealed the biosynthetic pathways of iP and ABA in red seaweeds similar to those in terrestrial plants. However, the mode of action in these seaweeds (Neopyropia yezoensis and Bangia fuscopurpurea) are dissimilar to those in terrestrial plants for IAA, iP, and ABA [162] and are yet to be investigated. Hormones like brassinosteroids (BRs), jasmonic acid, salicylic acid, and rhodomorphin have been reported from algae, but their possible signaling mechanism is yet to be investigated [54,141,163]. To date, their mode of action has not been elucidated. However, it has been proposed that their mode of action in algae could be similar to those of higher plants, which has been provided in Supplementary Materials.

7. Mode of Action of Pheromones in Algae

Green algae undergo sexual reproduction during their life cycle to survive unfavorable environmental conditions. Induction of gametogenesis in Chlamydomonas reinhardtii is activated via a reduced nitrogen supply in the environment [164]. After gametogenesis, the agglutinins, sex-specific glycoproteins located on the flagella, are synthesized, which promotes the interactions between different mating types and may lead to their fusion [165]. The gamete fusion takes place only when two compatible gametes come in contact with each other [164]. This contact induces the production of certain enzymes, which facilitate their fusion and form a cell having four flagella, ultimately giving rise to a non-motile zygote [96]. The attraction between the gametes and the level of compatibility between cells in C. reinhardtii decides the mating success. Similarly, in other species of Chlamydomonas, chemotactic behavior of gametes can be observed [74]. At low concentrations of pheromone lurlene, motile MT− gametes of Chlamydomonas allensworthii attract the motile MT+ gametes [166].

In Volvox carteri f. nagariensis, the male clones produce inducer molecules, extracellular matrix (ECM) glycoproteins called pherophorins, which can control sexualization. This protein is originally of somatic origin but can induce the production of respective gametes in both male and female algae [167,168]. After the gamete production, the sperm cells attain the ability to produce this inducer, which is now called pheromone [169]. The release of this pheromone can also lead to the production of hydroxyproline. Remarkably, the wounding of Volvox also produces a similar protein. Such expression of the same genes, which are activated by the wounding as well as pheromone induction, hints toward an existing relationship between environmental stress, sexual reproduction, and wound healing at the molecular level [170].

The blade cells of green macroalga Ulva mutabilis secrete some regulatory factors, which control the gametogenesis and are important to keep its thallus in a vegetative state. One of these factors, a sporulation inhibitor, prevents the differentiation of blade cells into gametangia [60]. During the maturation of the thallus, the production of this factor gradually decreases and stops until the concentration drops to the inhibitory concentration of 10⁻¹⁴ M. At this point, another sporulation inhibitor is released into the environment, which can supposedly control the distribution of gametangia spatially while they are developing. After induction, a swarming inhibitor is produced, which inhibits the release of gametes from U. mutabilis and U. lactuca [96].

Ectocarpene was identified as the first male-attracting chemical released by female plants. It was the first identified from Ectocarpus siliculosus. The compound is apolar and derived from a fatty acid, 9-hydroperoxyicosa (5Z,7E,11Z,14Z,17Z)-pentaenoic acid [61]. This compound is formed after the precursor is subjected to thermal rearrangement [104]. The inactivation process is wholly controlled by temperature and does not require any enzymatic activity [96]. Ectocarpene functions as a chemoattractant not only in many Ectocarpus species [62] but also in other brown algal genera like Adenocystis and Sphacelaria [63]. However, it has not yet been determined if the ectocarpene is as native in these species as in E. siliculosus. The gamete recognition and union in this alga is mediated by the lectin–glycoprotein complexes present in the membranes [171]. In another example, a structurally related epoxidized hydrocarbon from Laminaria digitata synchronizes the release of male gametes [172].

In marine brown algae Hormosira banksiii, Durvillea sp., Xiphophora sp., Scytosiphon lomentaria, and Colpomenia perergrina, chemical signal hormosirene was found to be released by female gametes (1–1000 pmol) to attract their conspecific male gametes [104]. Various life cycle stages of Giffordia mitchellae produce odoriferous compounds comprising mainly of giffordene and its stereoisomers because its male gametes are strongly attracted to settled female gametes [98]. In the orders Laminariales, Sporochnales, and Desmarestiales, sexual pheromones induce spermatozoid release from antheridia. In Laminaria, Maier et al. summarized the regulation of sexual reproduction by pheromones and other environmental factors [173]. The chemotactic movement of spermatozoids of Hormosira banksii and Laminaria digitata has also been reported in the literature [174,175]. Wirth and Boland recognized spermatozoid-attracting and spermatozoid-releasing factors in Perithalia caudata [176].

The interactions between various receptor–pheromone complexes have been studied in many species utilizing a number of pheromone analogs synthesized chemically [173]. But still, there seems to be no clarity on the molecular nature and cellular localization of various pheromone receptors. The first stage in the binding of brown algal pheromones is probably the partitioning into the cellular membrane due to the hydrophobic nature of this compound. Binding to the receptor protein strongly depends on the steric characteristics of the pheromone molecules and is intermediated by non-covalent dispersion forces, like how the double bonds are arranged in the molecule [74]. In chemotaxis assays, the binding process involves a strict enantiomer differentiation, which occurs as per the enantiomer specificity; the higher the specificity, the better the binding. Boland et al. hypothesized computer scheming for the identification of minimum energy conformations and a receptor-bound metal cation acting as the coordination center in pheromone binding [177].

8. Methods for Extraction, Identification, and Quantification of Hormones from Algae

A number of traditional extraction (with solvent) and novel extraction methods have been used to extract the phytohormones from plant samples. Of late, novel methods including microwave-assisted extraction (MAE), ultrasound-assisted extraction (UAE), pressurized liquid extraction (PLE), enzyme-assisted extraction (EAE), and supercritical fluid extraction (SFE) have been standardized for hormone extraction from algae [178,179]. From the analytical point of view, some of the most common and advantageous methods to extract phytohormones are liquid–liquid extraction (LLE), different types of liquid microextraction (LME), solid-phase extraction (SPE), or molecularly imprinted extraction (MIPE) [180]. Different protocols for the identification and quantification of phytohormones from algae are summarized in the flow chart (Supplementary Figure S2).

Seaweed sap, obtained mechanically by expelling water from fresh Kappaphycus alvarezii, is filtered with a nylon cloth (20–50 μm mesh size). Auxins, gibberellins, and cytokinins can then be extracted from sap using DEE, ethyl acetate, and n-butanol, respectively. Samples are kept overnight for solvent evaporation, and the pellets are then dissolved in 1–2 mL of HPLC-grade methanol. Phytohormones are then identified using MS/MS followed by quantification with ESI-MS, and quantification is validated against HPLC [32]. Among other techniques, auxin, cytokinin, and abscisic acid have been extracted from red algae (Porphyra, Gelidium, Gracilaria, and Hypnea), endogenous auxin can be extracted with cold phosphate buffer containing 0.02% sodium diethyldithiocarbamate, cytokinin can be extracted with ice-cold ethanol (70%), and abscisic acid can be extracted with ice-cold methanol: water: acetic acid (10:89:1, v/v) containing sodium diethyldithiocarbamate (400 µg g⁻¹ dwt). Subsequently, solid-phase extraction followed by immune affinity chromatography can be used for purification and analyzed with liquid chromatography–tandem mass spectrometry [37]. The simultaneous determination of broad classes of phytohormones in Monostroma and different species of Ulva (U. fasciata, U. lactuca, U. taeniata, and U. linza), using the extraction buffer methanol: water: formic acid (15:4:1) and subsequent purification with help of dispersive liquid–liquid microextraction (DLLME) [181], followed by the HPLC coupled to ultraviolet detector has also been reported [182].

A mixture of acetonitrile 80% (v/v) and acetic acid 1% (v/v) can also be used as an extraction solvent for the comprehensive quantification of phytohormones (auxin (IAA), N6-(∆2-isopentenyl) adenine (iP), abscisic acid (ABA), and salicylic acid) in red seaweeds (Bangia fuscopurpurea and Pyropia yezoensis). After extraction, the acetonitrile extract is purified with solid-phase extraction and analyzed with an LC-ESI-MS/MS system [183]. Simultaneous analysis methodology of nine phytohormones from Cladophora glomerata and Spirulina sp. was developed and optimized by Górka and Wieczorek. In this, the samples were extracted with the SFE-CO₂ extraction method with controlled condition of 500 bar pressure at 40 °C and analyzed using reversed-phase high-performance liquid chromatography (RP-HPLC) with a photodiode array detector (PDA) [180]. In another method, the aqueous extracts of dried Padina durvillaei and Ulva lactuca (1:10 w/v) are lyophilized. The lyophilized extract is then suspended in 80% methanol (MeOH) (1% acetic acid). This is followed by centrifugation, drying of supernatant, and dissolution in 1% (v/v) acetic acid. It is then passed consecutively through the reverse-phase column. Finally, the acidic hormones are eluted with MeOH and basic hormones with 60% MeOH with 5% aqueous ammonia. Dried final residues are dissolved in 5% (v/v) acetonitrile, 1% (v/v) MeOH, and 1% acetic acid. The hormones can then be quantified using ultra-high-performance liquid chromatography–mass spectrometry (UHPLC-MS) [184].

Reliable analytical techniques such as GC/MS and UPLC-MS/MS can also be used to identify phytohormones in microalgae (Chlorella minutissima) [185]. Gibberellins and brassinosteroids can be extracted from the different microalgae using 80% acetonitrile containing 5% formic acid as a solvent and analyzed with ultra performance chromatography–tandem mass spectrometry (UPLC-MS/MS) [186]. Endogenous hormones in Chlorella minutissima are extracted in cold 50 mM phosphate buffer (pH 7.0) with sodium diethyldithiocarbamate and analyzed using ultra-high-performance liquid chromatography (UPLC) equipped with an electrospray interface (ESI) [185,187].

9. Methods for Extraction, Identification, and Quantification of Pheromones from Algae

Volatile compounds like pheromones can be isolated using different methods, namely: CO₂ extraction method, cold trap condensation, head-space, and closed-loop-stripping method. Among all the methods, the closed-loop extraction method by Grob and Zurcher has proven to be the most efficient [188,189]. Different protocols for the identification and quantification of pheromones from algae are summarized in the flow chart (Supplementary Figure S3).

The closed-loop extraction method was used by Maier et al. [190] for the Lamoxirene extraction from the egg secretions in Laminariales and by Maier and Clayton [191] for the Hormosirene extraction from Hormosira banksii. Volatile compounds (chemoattractants) from the seaweed Ulva pertusa (now Ulva australis) can be extracted by distillation method with CH₂Cl₂ or Pentane used as solvent [192]. To extract the volatiles from microalgae, the cold trap condensation method is widely used, e.g., ectocarpene was obtained using this method [193]. In this method, a purified air stream is passed briefly through female gametes’ culture media or suspensions of Skeletonema costatum and Lithodesmium undulatum, which is then directed into a cold trap (−50 °C), containing 4–5 mL of n-pentane [194]. Trapped volatile compounds from Ectocarpus, Fucus, and Cutleria can then be adsorbed on an activated carbon bed of 1.5 mg and then desorbed with dichloromethane [71]. Finally, the eluates are purified using glass-capillary gas chromatography and analyzed with GC-MS [71,194]. Another method of extraction is where the cultures of the bloom-forming dinoflagellate Alexandrium tamarense have been subjected to suction filtration onto 45 mm GF/F filters for cell lysis, which causes the release of pheromone compounds from cells. The cell-free filtrate is then pumped through a 100 mg ENV + SPE column to trap the exudates. The eluate is concentrated in a centrifugal concentrator and resolved in methanol: water (80:20) having 20 mM ammonium acetate. Further eluted compounds are analyzed using Fourier transform ion cyclotron resonance (FT-ICR) MS. The stable isotope pattern (SIP), a novel algorithm, and an automated workflow can be used for comparison with theoretical isotope intensity profiles to produce the empirical formulae of the detected metabolites [195].

10. Perspectives

The primary objective of this review is to provide a comprehensive account of hormones and pheromones from an algal point of view. This article can form the basis for benchmarking future investigations. A lot of parallels could be drawn between the higher plants and the algae, e.g., exposure to rapid climate changes, seasonal growth cycles, and biotic stress from herbivores. This and similar factors underline the fact that algal growth and development, like in higher plants, are influenced and controlled by hormones and pheromones. Similarly, hormones have a major role to play in defense mechanisms in higher angiospermic plants, and this could be said about algae as well but its molecular mechanism in algae has not been established thus far. Deciphering this mechanism could be a game changer in technological interventions in the cultivation of various algae, which are frequently infested by epi/endophytes. A lot of focus is being put on understanding the reproduction of various commercially important species, which could revolutionize the spore-based commercial cultivation technologies, and the study of hormones and pheromones might hold the key for future development. Recent studies have unequivocally provided evidence for the use of algal-based bio-stimulants in micro-algal and seaweed cultivation to enhance circular blue economy [196,197].

The applications in terms of biotechnological implications for the commercial aquaculture industry include induction of reproduction, germplasm enhancement, micro-propagation, and seedling production. The commercial farming of seaweeds—where clonal propagation is not practiced—overwhelmingly depends on the availability of reproductive cells, e.g., zoospores, gametes for seeding [Ulva, Porphyra (Pyropia), Undaria, etc.]. The use of hormones and pheromones can ascertain and ensure the timely availability of carpospores, tetraspores, zoospores, and gametes for seeding. Research has shown that treating the thalli of Grateloupia imbricata with methyl jasmonate reported a ~7.5-fold increase in cystocarp numbers. Further, the maturation occurred within 48 h of treatment, thereby shortening the typical >3-week maturation period [198]. Similarly, the use of ethylene as a physiological regulator in the tetrasporogenesis of Pterocladiella capillacea has been established [199]. The thalli exposed to ethylene for 15 and 30 min produced a high number of tretrasporangia. Polyamines like spermidine and spermine were reported to enhance cystocarp maturation followed by spore release in Gracilaria cornea [52]. The exogenous application of ABA to Saccharina japonica enhanced the formation of sporophytic sori [125]. These studies unequivocally confirmed the role and thus potential role of plant hormones in spore-mediated aquaculture of economically important seaweeds.

Another application is their use in germplasm enhancement. Studies have shown that the use of ethylene precursor, which is 1-aminocylopropane-1-carboxylic acid (ACC), provides Neopyropia yezoensis gametophytes tolerance to heat stress [126,127,200]. It was shown that Pyropia orbicularis had 4–7 times higher free ABA levels during water deficit conditions [201]. Further, ABA is known to regulate the activation of the antioxidant enzymes during desiccation. The long distance transportation of planting material in red seaweed Kappaphycus alvarezii reported low survival and vigor. Thus, exogenous application of ABA to these seedlings could provide desiccation stress tolerance for germplasm improvement.

The role of plant hormones in micro-propagation and seedling production is well known. The use of indole-3-acetic acid, 2,4-dichlorophenoxyacetic acid (2,4-D), and kinetin on callus formation, growth, and regeneration of Gracilaria tenuistipitata and G. perplexa was reported [202]. Similarly, auxins and cytokinins are known to help the tissue culture of Grateloupia dichotoma [112]. The plant growth hormones like 2,4-D, IAA, and α-naphthalene acetic acid were found to be effective in successful micro-propagation for the production of clonal planting materials in Gracilaria changii and Kappaphycus alvarezii through tissue culture [203]. The plant hormones individually as well as in combination have reported improved survival and regeneration in Gracilaria dura [204].

Therefore, the understating of endogenous level and exogenous application have several pivotal implications in seaweed aquaculture. Unlike higher plants, the application domain for plant hormones and pheromones has yet to expand for seaweeds. Decreasing genetic variability of seaweeds due to repeated conventional vegetative propagation methods in seaweed cultivation causes decreased growth rates and productivity and makes them more susceptible to diseases [205]. Micro-propagation through tissue culture has been an alternative to this conventional method. Plant growth regulators and their role in the tissue culture of marine macroalgae (seaweeds) have been extensively reviewed [21,111,206], suggesting that the phytohormones play an important role in the growth and development of algae. Further, significant efforts are needed to better understand the physiological effects and varied functions of hormones, which depend on the development of different groups of algal taxonomy. Male-sterile Gelidium vagum macroalgae exhibited notably faster growth rates compared to plants from the different lines, indicating that male-sterile gametophytes might be more suitable for aquaculture than the typical wild-type plants of this species [207].

Exogenous application of α-naphthaleneacetic acid (NAA), gibberellin (GA3), and 6-benzyl adenine (BAP) has been used to induce regeneration of Sargassum fusiforme to obtain more regenerative seedlings within a short time, thereby improving economic efficiency [208]. Plant growth hormone (zeatin + phenylacetic acid) has been used for micro-propagation in three color morphotypes of Kappaphycus alvarezii var. “adik-adik” [209]. The successful regeneration of young plants from various Kappaphycus varieties was achieved using a culture medium composed of zeatin and phenylacetic acid (PAA) along with Ascophyllum/Acadian marine plant extract powder (AMPEP) obtained from the brown seaweed Ascophyllum nodosum [210]. AMPEP concentrations with 0.1, 1.0 mg L⁻¹ stimulate the growth in macroalgae Gracilaria caudata and Gracilaria corticata var. cylindrica, and at higher concentrations of 5.0 mg L⁻¹, it stimulates the growth in Laurencia catarinensis [211,212]. Seaweed extracts containing hormones (auxins, gibberellins, cytokinins, gibberellins, abscisic acid, ethylene, betaine, and polyamines) have more potential as bio-stimulants in agriculture [213]. Phytohormones’ abscisic acid (ABA), 24-epibrassinolide (EBL), brassinolide (BL), and indole-3-acetic acid (IAA) showed the stimulatory effects on microalga Scenedesmus quadricauda cell growth, biomass production, intracellular concentrations of chlorophyll-a, carotenoid, and lipids biosynthesis [214]. Phytohormones in combination are widely used to enhance algal cultivation. In Chlorella sorokiniana, combined treatments of NAA_5ppm + GA3_10ppm + zeatin_1ppm increased the biomass production by 170% as compared to control, followed by treatments of NAA_5ppm + GA3_10ppm and NAA_2.5ppm + zeatin_1ppm stimulating the biomass production by 138% and 136%, respectively [215]. Similarly, in freshwater microalgae, the combination of phytohormones has been reported to stimulate biomass and lipid production [216]. Pheromones can be used to facilitate the selective release of male and female gametes in marine macroalgae by controlling the release and response to specific pheromones [217,218]. Breeders can choose the desired individuals for mating, leading to the development of seaweed strains with desirable traits. Sexual signaling in marine benthic diatom Seminavis robusta is controlled by a complex and unknown pheromone system called sex-inducing pheromone (SIP+), which activates the shift from mitosis to meiosis in the opposing mating type. Additionally, it stimulates the activation of genes related to proline biosynthesis and the release of an attraction pheromone derived from proline [219].

In seaweed, gene regulation can occur in response to changes in environmental conditions, including hormone signaling. For example, the levels of certain genes related to growth or stress response can be upregulated or down-regulated in response to hormonal signals [141,220]. Stress hormone ABA increases in concentration in response to abiotic stress, initiating a range of physiological processes aimed at enhancing their survival chances [221]. In filamentous green alga Stigeocloflium cf. tenue, exogenous application of ABA (10⁻⁷ to 10⁻⁵ M) in a culture medium for the 3 weeks reported a slight reduction in growth and senescence. So the physiological concentrations of ABA below 10⁻⁷ to 10⁻⁵ M show tolerance and healthy growth, whereas concentrations above this for a longer period (3 weeks) may start growth reduction and senescence in Stigeocloflium cf. tenue [38]. Also, the endogenous ABA was reported to increase (10×) in microalga Draparnaldia mutabilis and JA increased (15×) in Draparnaldia salina when grown in osmotic stress (50 and 85 mM NaCl), suggesting that the endogenous ABA and JA play important roles in salt tolerance in microalgae [38]. In seaweeds, CRISPR technology can be used to target and modify genes involved in hormone production or response pathways in seaweed and shed light on the regulatory mechanisms [222].

It is worth mentioning that the field of seaweed hormone and pheromone research is relatively young compared to terrestrial plants, and there is ongoing research to better understand the role of hormones and pheromones in seaweed growth, health, and defense mechanisms. While these studies provide some insights, further research is needed to fully explore the potential of hormone and pheromone calibration for identifying elite, healthy, and disease-free seaweed germplasm.

11. Way Forward

It is well evident that plant hormones play a pivotal role in regulating growth and development while pheromones are needed for induction of reproduction and sexual maturity in seaweeds. Although plant hormones possess specific functions, they tend to mediate with other hormones either antagonistically or cooperatively via complex crosstalk for achieving optimal outcomes, and their mode of action in isolation is most unlikely in nature.

The present review for the first time provides types and diversity of biosynthetic pathways of five phytohormones and five pheromones reported in algal representatives (along with six hormones and two pheromones in higher angiospermic plants but possibly also operational in algal representatives); modes of action of five hormones and five pheromones reported in algal representatives (along with the modes of action of five hormones in higher angiospermic plants but possibly also operational in algal representatives); roles of 11 hormones and 29 pheromones reported in algal representatives; and extraction protocols of four hormones and six pheromones reported in algal representatives. The application of endogenous plant hormones as environmentally friendly or organic bio-stimulants for higher crops and seaweeds is well documented [196]. It has been shown that in higher plants, hormones, besides their usual known functions, are also involved in the interactions between them and beneficial microbes [223]. Further, these microbial-derived hormones assist in imparting tolerance to biotic and abiotic stress in higher angiospermic plants. However, such studies are seldom attempted in seaweeds and are necessary so that such knowledge can be applied to sustainable seaweed aquaculture. Similarly, the use of plant hormones for alleviating stress tolerance and disease resistance would be critical to retard the aggravated impacts of climate change in terms of mass mortality and failure of commercial seaweed crops. The diversity in sexual strategies in seaweeds provides ample opportunity in the presence of chemical diversity of pheromones, but the research in elucidating functional roles is slow. The successful application of pheromones in triggering sexual reproduction and life cycle transition in Ulva has been evident [60,144]. The microbial-derived chemicals are also known to enhance reproduction in green and red seaweeds [224]. Unlike plant hormones, analogs for pheromones are not widely studied. This is essentially due to the minuscule amount released into the environment, the absence of standard extraction procedures, and a vast range of active chemical moieties, e.g., non-polar small hydrocarbons to polar glycoproteins of high molecular weight with respect to pheromones. Further studies in this direction would help elicit physiological responses reminiscent of pheromones to aid control over reproduction; successful breeding; and continuous and sustainable seedling production by removing seasonality barriers to enhance aquaculture prospects.

The exact identification and quantification of plant hormones and pheromones from seaweeds have been usually performed using ultra-high-performance liquid chromatography–mass spectrometry methodology. The current technical advancements in methodologies have focused on developing protocols that are less labor-intensive and comprehensive rather than improving sensitivity and accuracy [162]. However, fine quantification is difficult to conduct due to the interfering effects of cellular constituents. Novel methods in extraction and further identification and quantifications are needed for quick and reliable outcomes. Consequently, sophisticated imaging tool kits applied with a fluorescence-compatible clearing approach with synthetic transcriptional reporters for specific plant hormones or pheromones in seaweeds may be used for determining the precise spatial and temporal regulation. The implementation of such cutting-edge techniques would open new possibilities of enhancing our ability to understand the mode of action and characterize hormone and pheromone dynamics at the cellular level. In the high-throughput-mediated multi-omics era, the understating of the mechanism of hormones and pheromones at the metabolomics, proteomics, transcriptomics, and genomics levels would shed more light not only on their biosynthesis, mode of action, and signal transduction but also in a micro-evolutionary context. There seems enormous potential for involving multidisciplinary teams including bioinformatics promoting a paradigm shift in our understating.

Supplementary Materials

The following supporting information can be downloaded at: https://0-www-mdpi-com.brum.beds.ac.uk/article/10.3390/phycology4010001/s1, The Biosynthetic pathways and mode of actions in higher angiospermic plants are given in the Supplementary Information, Supplementary Figure S1, and Supplementary Figures S4–S10. References [225,226,227,228,229,230,231,232,233,234,235,236,237,238,239,240,241,242,243,244,245,246,247,248,249,250,251,252,253,254,255,256,257] are cited in the Supplementary Materials. Figure S1: Tryptophan-independent pathway of auxin biosynthesis in higher plants., Figure S2: Flow chart summarizing the protocols for identification and quantification of phytohormones from algae., Figure S3. Flow chart summarizing the protocols for identification and quantification of pheromones from algae., Figure S4. Biosynthesis of brassinosteroids in higher plants., Figure S5: Biosynthesis of jasmonic acid (JA) in higher plants., Figure S6: Biosynthesis of polyamines in higher plants., Figure S7: Biosynthesis of salicylic acid (SA) in higher plants., Figure S8: Biosynthesis of strigolactones (SL) in higher plants., Figure S9: Biosynthesis of finavarrene from dodeca-3,6,9-trienoic acid in higher plants., Figure S10: Biosynthesis of Hormosirene in higher plants.

Author Contributions

Conceptualization, V.A.M.; data curation, S.G.R. and S.B.; writing—original draft preparation, S.G.R., S.B. and V.A.M.; visualization, S.G.R., S.B. and V.A.M.; writing—review and editing, S.G.R., S.B. and V.A.M.; supervision, V.A.M.; project administration, V.A.M.; funding acquisition, V.A.M. All authors have read and agreed to the published version of the manuscript.

Funding

SGR thank University Grant Commission, New Delhi, for the award of Senior Research Fellowship [817/(CSIR-UGC NET DEC.2018)]. Authors would like to thank Council for Scientific and Industrial Research, New Delhi, for funding (Project No. HCP-0024).

Data Availability Statement

The data-sets generated during and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Acknowledgments

We are thankful to Director, CSIR-CSMCRI for encouragement and facilities. The authors thank Rahul D. Patil and Shivprasad S. Patil for help during manuscript preparation. This manuscript has PRIS registration number 101/2022.

Conflicts of Interest

The authors declare no conflict of interest.

References

Hoek, C.; Mann, D.; Jahns, H.M. Algae: An Introduction to Phycology; Cambridge University Press: Cambridge, UK, 1995. [Google Scholar]
Koushalya, S.; Vishwakarma, R.; Malik, A. Unraveling the diversity of algae and its biomacromolecules. Microb. Nat. Macromol. 2021, 1, 179–204. [Google Scholar] [CrossRef]
Rajauria, G.; Cornish, L.; Ometto, F.; Msuya, F.E.; Villa, R. Identification and Selection of Algae for Food, Feed, and Fuel Applications. In Seaweed Sustainability: Food and Non-Food Applications; Academic Press: New York, NY, USA, 2015; pp. 315–345. [Google Scholar] [CrossRef]
Dittami, S.M.; Heesch, S.; Olsen, J.L.; Collén, J. Transitions between marine and freshwater environments provide new clues about the origins of multicellular plants and algae. J. Phycol. 2017, 53, 731–745. [Google Scholar] [CrossRef] [PubMed]
Leliaert, F.; Verbruggen, H.; Zechman, F.W. In to the deep: New discoveries at the base of the green plant phylogeny. Bioessays 2011, 33, 683–692. [Google Scholar] [CrossRef] [PubMed]
Chung, I.K.; Beardall, J.; Mehta, S.; Sahoo, D.; Stojkovic, S. Using marine macroalgae for carbon sequestration: A critical appraisal. J. Appl. Phycol. 2011, 23, 877–886. [Google Scholar] [CrossRef]
Druehl, L. Mouritsen, O.G. 2013. Seaweeds, Edible, Available & Sustainable. University of Chicago Press, 283 pp. ISBN 978-0-226-04436-1. J. Phycol. 2013, 49, 1229. [Google Scholar] [CrossRef]
Cai, J.; Lovatelli, A.; Aguilar-Manjarrez, J.; Cornish, L.; Dabbadie, L.; Desrochers, A.; Diffey, S.; Garrido Gamarro, E.; Geehan, J.; Hurtado, A.; et al. Seaweeds and Microalgae: An Overview for Unlocking Their Potential in Global Aquaculture Development. In FAO Fisheries and Aquaculture Circular; FAO: Rome, Italy, 2021; pp. 1–36. [Google Scholar] [CrossRef]
Hurtado, A.Q. Genetic Resources for Farmed Seaweeds. In Thematic Background Study; FAO: Rome, Italy, 2022. [Google Scholar] [CrossRef]
González-Gloria, K.D.; Rodríguez-Jasso, R.M.; Aparicio, E.; González, M.L.C.; Kostas, E.T.; Ruiz, H.A. Macroalgal biomass in terms of third-generation biorefinery concept: Current status and techno-economic analysis–A review. Bioresour. Technol. 2021, 16, 100863. [Google Scholar] [CrossRef]
Duerte, C.M.; Bruhn, A.; Krause-Jensen, D. A seaweed aquaculture imperative to meet global sustainability targets. Nat. Sustain. 2022, 5, 185–193. [Google Scholar] [CrossRef]
McHugh, D.J. A Guide to the Seaweed Industry; Food and Agriculture Organization of the United Nations: Rome, Italy, 2003; Volume 441, p. 105. [Google Scholar]
Khan, W.; Rayirath, U.P.; Subramanian, S.; Jithesh, M.N.; Rayorath, P.; Hodges, D.M.; Critchley, A.T.; Craigie, J.S.; Norrie, J.; Prithiviraj, B. Seaweed extracts as biostimulants of plant growth and development. J. Plant Growth Regul. 2009, 28, 386–399. [Google Scholar] [CrossRef]
Verkleij, F.N. Seaweed extracts in agriculture and horticulture: A review. Biol. Agric. Hortic. 1992, 8, 309–324. [Google Scholar] [CrossRef]
Karthikeyan, K.; Shanmugam, M. The effect of potassium-rich biostimulant from seaweed Kappaphycus alvarezii on yield and quality of cane and cane juice of sugarcane var. Co 86032 under plantation and ratoon crops. J. Appl. Phycol. 2017, 29, 3245–3252. [Google Scholar] [CrossRef]
Agarwal, P.K.; Dangariya, M.; Agarwal, P. Seaweed extracts: Potential biodegradable, environmentally friendly resources for regulating plant defence. Algal Res. 2021, 58, 102363. [Google Scholar] [CrossRef]
Trivedi, K.; Vijay Anand, K.G.; Kubavat, D.; Patidar, R.; Ghosh, A. Drought alleviatory potential of Kappaphycus seaweed extract and the role of the quaternary ammonium compounds as its constituents towards imparting drought tolerance in Zea mays L. J. Appl. Phycol. 2018, 30, 2001–2015. [Google Scholar] [CrossRef]
Crouch, I.J.; Van Staden, J. Effect of seaweed concentrate from Ecklonia maxima (Osbeck) Papenfuss on Meloidogyne incognita infestation on tomato. J. Appl. Phycol. 1993, 5, 37–43. [Google Scholar] [CrossRef]
Banu, A.T.; Ramani, P.S.; Murugan, A. Effect of seaweed coating on quality characteristics and shelf life of tomato (Lycopersicon esculentum mill). Food Sci. Hum. Wellness 2020, 9, 176–183. [Google Scholar] [CrossRef]
Mlambo, P.Z. Exploring the Fertiliser Potential of Biosolids from Algae Integrated Wastewater Treatment Systems. Ph.D. Thesis, Rhodes University, Grahamstown, South Africa, October 2013. [Google Scholar]
Bradley, P.M. Plant hormones do have a role in controlling growth and development of algae. J. Phycol. 1991, 27, 317–321. [Google Scholar] [CrossRef]
Singh, G.; Patel, A.; Tiwari, S.; Gupta, D.; Prasad, S.M. Signaling molecules hydrogen sulfide (H₂S) and nitric oxide (NO): Role in microalgae under adverse environmental conditions. Acta Physiol. Plant. 2022, 44, 68. [Google Scholar] [CrossRef]
Delaux, P.M.; Xie, X.; Timme, R.E.; Puech-Pages, V.; Dunand, C.; Lecompte, E.; Delwiche, C.F.; Yoneyama, K.; Bécard, G.; Séjalon-Delmas, N. Origin of strigolactones in the green lineage. New Phytol. 2012, 195, 857–871. [Google Scholar] [CrossRef] [PubMed]
Fries, L.; Aberg, S. Morphogenetic effects of phenylacetic and p-OH-phenylacetic acid on the green alga Enteromorpha compressa (L.) Grev. in axenic culture. J. Plant Physiol. 1978, 88, 383–388. [Google Scholar] [CrossRef]
Zhang, W.; Yamane, H.; Chapman, D.J. The phytohormone profile of the red alga Porphyra perforata. Bot. Mar. 1993, 36, 257–266. [Google Scholar] [CrossRef]
Provasoli, L. Effect of plant hormones on Ulva. Biol. Bull. 1958, 114, 375–384. [Google Scholar] [CrossRef]
Imahori, K.; Iwasa, K. Pure culture and chemical regulation of the growth of charophytes. Phycologia 1965, 4, 127–134. [Google Scholar] [CrossRef]
Jacobs, W.P. Are angiosperm hormones present in, and used as hormones by, algae? In Proceedings of the 12th International Conference on Plant Growth Substances; Bopp, M., Ed.; Springer: Berlin/Heidelberg, Germany, 1986; pp. 249–256. [Google Scholar] [CrossRef]
Mowat, J.A. A survey of results on the occurrence of auxins and gibberellins in algae. Bot. Mar. 1965, 8, 149–155. [Google Scholar] [CrossRef]
Stirk, W.A.; Novak, O.; Strnad, M.; van Staden, J. Cytokinins in macroalgae. Plant Growth Regul. 2003, 41, 13–24. [Google Scholar] [CrossRef]
Dworetzky, B.; Klein, R.M.; Cook, P.W. Effect of growth substances on “Apical Dominance” in Sphacelaria furcigera (Phaeophyta). J. Phycol. 1980, 16, 239–242. [Google Scholar] [CrossRef]
Prasad, K.; Das, A.K.; Oza, M.D.; Brahmbhatt, H.; Siddhanta, A.K.; Meena, R.; Eswaran, K.; Rajyaguru, M.R.; Ghosh, P.K. Detection and quantification of some plant growth regulators in a seaweed-based foliar spray employing a mass spectrometric technique sans chromatographic separation. J. Agric. Food Chem. 2010, 58, 4594–4601. [Google Scholar] [CrossRef] [PubMed]
Zhang, W.; Chapman, D.J.; Phinney, B.O.; Spray, C.R.; Yamane, H.; Takahashi, N. Identification of Cytokinins in Sargassum muticum (Phaeophyta) and Porphyra perforata (Rhodophyta). J. Phycol. 1991, 27, 87–91. [Google Scholar] [CrossRef]
Borowczak, E.; Kentzer, T.; Potulska-Klein, B. Effect of gibberellin and kinetin on the regeneration ability of Fucus vesiculosus L. Biol. Plant. 1977, 19, 405–412. [Google Scholar] [CrossRef]
Kwiatkowska, M.; Godlewski, M. Studies on the role of gibberellins in the regulation of spermatogenesis in Chara vulgaris L. Acta Soc. Bot. Pol. 1988, 57, 547–553. [Google Scholar] [CrossRef]
Stirk, W.A.; Novak, O.; Hradecka, V.; Pencík, A.; Rolcík, J.; Strnad, M.; van Staden, J. Endogenous cytokinins, auxins and abscisic acid in Ulva fasciata (Chlorophyta) and Dictyota humifusa (Phaeophyta): Towards understanding their biosynthesis and homoeostasis. Eur. J. Phycol. 2009, 44, 231–240. [Google Scholar] [CrossRef]
Yokoya, N.S.; Stirk, W.A.; van Staden, J.; Novák, O.; Turečková, V.; Pěnčí, K.A.; Strnad, M. Endogenous cytokinins, auxins, and abscisic acid in red algae from Brazil. J. Phycol. 2010, 46, 1198–1205. [Google Scholar] [CrossRef]
Tietz, A.; Ruttkowski, U.; Kohler, R.; Kasprik, W. Further investigations on the occurrence and the effects of abscisic acid in algae. Biochem. Physiol. Pflanz. 1989, 184, 259–266. [Google Scholar] [CrossRef]
Waaland, S.D. Hormonal Coordination of the Processes Leading to Cell Fusion in Algae: A Glycoprotein Hormone from Red Algae. In Proceedings of the 12th International Conference on Plant Growth Substances, Berlin/Heidelberg, Germany, 26–31 August 1985; Bopp, M., Ed.; Springer: Berlin/Heidelberg, Germany, 1986; pp. 257–262. [Google Scholar] [CrossRef]
Starr, R.C.; Jaenicke, L. Purification and characterization of the hormone initiating sexual morphogenesis in Volvox carteri f. nagariensis Iyengar. Proc. Natl. Acad. Sci. USA 1974, 71, 1050–1054. [Google Scholar] [CrossRef] [PubMed]
Driessche, T.V.; Jerebzoff, S.; Jerebzoff-Quintin, S. Phase-shifting effects of indole-3-acetic acid and the synchronization of three circadian metabolic rhythms in Acetabularia. Biol. Rhythm Res. 1988, 19, 81–87. [Google Scholar] [CrossRef]
Stirk, W.A.; Tarkowska, D.; Turecova, V.; Strnad, M.; Van Staden, J. Abscisic acid, gibberellins and brassinosteroids in Kelpak^®, a commercial seaweed extract made from Ecklonia maxima. J. Appl. Phycol. 2014, 26, 561–567. [Google Scholar] [CrossRef]
Christov, C.; Pouneva, I.; Bozhkova, M.; Toncheva, T.; Fournadzieva, S.; Zafirova, T. Influence of temperature and methyl jasmonate on Scenedesmus incrassulatus. Biol. Plant. 2001, 44, 367–371. [Google Scholar] [CrossRef]
Bouarab, K.; Adas, F.; Gaquerel, E.; Kloareg, B.; Salaun, J.P.; Potin, P. The innate immunity of a marine red alga involves oxylipins from both the eicosanoid and octadecanoid pathways. Plant Physiol. 2004, 135, 838–1848. [Google Scholar] [CrossRef] [PubMed]
Cohen, E.; Arad, S.; Heimer, Y.H.; Mizrahi, Y. Polyamine biosynthetic enzymes in the cell cycle of Chlorella: Correlation between ornithine decarboxylase and DNA synthesis at different light intensities. Plant Physiol. 1984, 74, 385–388. [Google Scholar] [CrossRef] [PubMed]
Theiss, C.; Bohley, P.; Voigt, J. Regulation by polyamines of ornithine decarboxylase activity and cell division in the unicellular green alga Chlamydomonas reinhardtii. Plant Physiol. 2002, 128, 1470–1479. [Google Scholar] [CrossRef]
Lee, T.M. Investigations of some intertidal green macroalgae to hyposaline stress: Detrimental role of putrescine under extreme hyposaline conditions. Plant Sci. 1998, 138, 1–8. [Google Scholar] [CrossRef]
Garcia-Jimenez, P.; Just, P.M.; Delgado, A.M.; Robaina, R.R. Transglutaminase activity decrease during acclimation to hyposaline conditions in marine seaweed Grateloupia doryphora (Rhodophyta, Halymeniaceae). J. Plant Physiol. 2007, 164, 367–370. [Google Scholar] [CrossRef]
Garcia-Jimenez, P.; Rodrigo, M.; Robaina, R.R. Influence of plant growth regulators, polyamines and glycerol interaction on growth and morphogenesis of carposporelings of Grateloupia cultured in vitro. J. Appl. Phycol. 1998, 10, 95–100. [Google Scholar] [CrossRef]
Marián, F.D.; García-Jiménez, P.; Robaina, R.R. Polyamines in marine macroalgae: Levels of putrescine, spermidine and spermine in the thalli and changes in their concentration during glycerol-induced cell growth in vitro. Physiol. Plant. 2000, 110, 530–534. [Google Scholar]
Sacramento, A.T.; García-Jimenez, P.; Robaina, R.R. The polyamine spermine induces cystocarp development in the seaweed Grateloupia (Rhodophyta). Plant Growth Regul. 2007, 53, 147–154. [Google Scholar] [CrossRef]
Guzman-Uriostegui, A.; García-Jimenez, P.; Marian, F.D.; Robledo, D.; Robaina, R.R. Polyamines influence maturation in reproductive structures of Gracilaria cornea (Gracilariales, Rhodophyta). J. Phycol. 2002, 38, 1169–1175. [Google Scholar] [CrossRef]
Sacramento, A.T.; Garcia-Jimenez, P.; Alcazar, R.; Tiburcio, A.; Robaina, R.R. Influence of polyamines on the sporulation of Grateloupia (Halymeniaceae, Rhodophyta). J. Phycol. 2004, 50, 887–894. [Google Scholar] [CrossRef]
Zhou, B.; Tang, X.; Wang, Y. Salicylic acid and heat acclimation pretreatment protects Laminaria japonica sporophyte (Phaeophyceae) from heat stress. Chin. J. Oceanol. Limnol. 2010, 28, 924–932. [Google Scholar] [CrossRef]
Jennings, R.C. Gibberellins as endogenous growth regulators in green and brown algae. Planta 1968, 80, 34–42. [Google Scholar] [CrossRef]
Abe, H.; Marumo, S. Identification of auxin-active substances as ethyl chlorogenate and indolyl-3-acetic acid in immature seeds of Helianthus annuus. Agric. Biol. Chem. 1972, 36, 42–46. [Google Scholar] [CrossRef]
Brain, K.R.; Chalopin, M.C.; Turner, T.D.; Blunden, G.; Wildgoose, P.B. Cytokinin activity of commercial aqueous seaweed extract. Plant Sci. Lett. 1973, 1, 241–245. [Google Scholar] [CrossRef]
Basu, S.; Sun, H.; Brian, L.; Quatrano, R.L.; Muday, G.K. Early embryo development in Fucus distichus is auxin sensitive. Plant Physiol. 2002, 130, 292–302. [Google Scholar] [CrossRef]
Karlson, P.; Lüscher, M. ‘Pheromones’: A new term for a class of biologically active substances. Nature 1959, 183, 55–56. [Google Scholar] [CrossRef]
Stratmann, J.; Paputsoglu, G.; Oertel, W. Differentiation of Ulva mutabilis (Chlorophyta) gametangia and gamete release are controlled by extracellular inhibitors. J. Phycol. 1996, 32, 1009–1021. [Google Scholar] [CrossRef]
Muller, D.G.; Jaenicke, L.; Donike, M.; Akintobi, T. Sex attractant in a brown alga: Chemical structure. Science 1971, 171, 815–817. [Google Scholar] [CrossRef] [PubMed]
Muller, D.G.; Gassmann, G. Sexual hormone specificity in Ectocarpus and Laminaria (Phaeophyceae). Nat. Sci. 1980, 67, 462–463. [Google Scholar] [CrossRef]
Muller, D.G.; Boland, W.; Jaenicke, L.; Gassmann, G. Diversification of chemoreceptors in Ectocarpus, Sphacelaria, and Adenocystis (Phaeophyceae). J. Nat. Sci. 1985, 40, 457–459. [Google Scholar] [CrossRef]
Hay, M.E.; Piel, J.; Boland, W.; Schnitzler, I. Seaweed sex pheromones and their degradation products frequently suppress amphipod feeding but rarely suppress sea urchin feeding. Chemoecology 1998, 8, 91–98. [Google Scholar] [CrossRef]
Phillips, J.A.; Clayton, M.N.; Maier, I.; Boland, W.; Muller, D.G. Sexual reproduction in Dictyota diemensis (Dictyotales, Phaeophyta). Phycologia 1990, 29, 367–379. [Google Scholar] [CrossRef]
Hay, M.E.; Duffy, J.E.; Pfister, C.A.; Fenical, W. Chemical defense against different marine herbivores: Are amphipods insect equivalents? Ecology 1987, 68, 1567–1580. [Google Scholar] [CrossRef]
Hay, M.E.; Fenical, W. Marine plant-herbivore interactions: The ecology of chemical defense. Annu. Rev. Ecol. Evol. Syst. 1988, 19, 111–145. [Google Scholar] [CrossRef]
Schnitzler, I.; Boland, W.; Hay, M.E. Organic sulfur compounds from Dictyopteris spp. deter feeding by an herbivorous amphipod (Ampithoe longimana) but not by an herbivorous sea urchin (Arbacia punctulata). J. Chem. Ecol. 1998, 24, 1715–1732. [Google Scholar] [CrossRef]
Maier, I.; Muller, D.G.; Gassmann, G.; Boland, W.; Marner, F.J.; Jaenicke, L. Pheromone-triggered gamete release in Chorda tomentosa. Nat. Sci. 1984, 71, 48–49. [Google Scholar] [CrossRef]
Muller, D.G.; Jaenicke, L. Fucoserraten, the female sex attractant of Fucus serratus L. (Phaeophyta). FEBS Lett. 1973, 30, 137–139. [Google Scholar] [CrossRef] [PubMed]
Maier, I.; Muller, D.G. Sexual pheromones in algae. Biol. Bull. 1986, 170, 145–175. [Google Scholar] [CrossRef]
Jaenicke, L. One hundred and one years of chemotaxis. Pfeffer, pheromones, and fertilization. Bot. Acta 1988, 101, 149–159. [Google Scholar] [CrossRef]
Muller, D.G. The role of pheromones in sexual reproduction of brown algae. Plant Biol. 1989, 7, 201–213. [Google Scholar]
Maier, I. Gamete orientation and induction of gametogenesis by pheromones in algae and plants. Plant Cell Environ. 1993, 16, 891–907. [Google Scholar] [CrossRef]
Jaenicke, L.; Muellar, D.G.; Moore, R.E. Multifidene and aucantene, C11 hydrocarbons in the male-attracting essential oil from the gynogametes of Cutleria multifida (Phaeophyta). J. Am. Chem. Soc. 1974, 96, 3324–3325. [Google Scholar] [CrossRef]
McConnell, O.J.; Fenical, W. Ochtodene and ochtodiol: Novel polyhalogenated cyclic monoterpenes from the red seaweed Ochtodes secundiramea. J. Org. Chem. 1978, 43, 4238–4241. [Google Scholar] [CrossRef]
Chandler, J.W. Local auxin production: A small contribution to a big field. Bioessays 2009, 31, 60–70. [Google Scholar] [CrossRef]
Kiseleva, A.A.; Tarachovskaya, E.R.; Shishova, M.F. Biosynthesis of phytohormones in algae. J. Plant Physiol. 2012, 59, 595–610. [Google Scholar] [CrossRef]
Zhao, Y. Auxin biosynthesis and its role in plant development. Annu. Rev. Plant Biol. 2010, 61, 49–64. [Google Scholar] [CrossRef] [PubMed]
Mok, M.C.; Martin, R.C.; Mok, D.W. Cytokinins: Biosynthesis metabolism and perception. Vitr. Cell. Dev. Biol. Plant 2000, 36, 102–107. [Google Scholar] [CrossRef]
Lindner, A.C.; Lang, D.; Seifert, M.; Podlešáková, K.; Novák, O.; Strnad, M.; Reski, R.; von Schwartzenberg, K. Isopentenyltransferase-1 (IPT1) knockout in Physcomitrella together with phylogenetic analyses of IPTs provide insights into evolution of plant cytokinin biosynthesis. J. Exp. Bot. 2014, 65, 2533–2543. [Google Scholar] [CrossRef] [PubMed]
Ordog, V.; Stirk, W.A.; van Staden, J.; Novak, O.; Strnad, M. Endogenous cytokinins in three genera of microalgae from the Chlorophyta. J. Phycol. 2004, 40, 88–95. [Google Scholar] [CrossRef]
Von Schwartzenberg, K.; Nunez, M.F.; Blaschke, H.; Dobrev, P.I.; Novak, O.; Motyka, V.; Strnad, M. Cytokinins in the Bryophyte Physcomitrella patens: Analysis of activity, distribution, and cytokinin oxidase/dehydrogenase overexpression reveal the role of extracellular cytokinins. Plant Physiol. 2007, 145, 786–800. [Google Scholar] [CrossRef]
Taiz, L.; Zeiger, E.; Møller, I.M.; Murphy, A. Plant Physiology and Development, 6th ed.; Sinauer Associates Incorporated: Sunderland, UK, 2015; p. 761. [Google Scholar]
Schwender, J.; Seemann, M.; Lichtenthaler, H.K.; Rohmer, M. Biosynthesis of isoprenoids (carotenoids, sterols, prenyl side-chains of chlorophylls and plastoquinone) via a novel pyruvate/glyceraldehyde 3-phosphate non-mevalonate pathway in the green alga Scenedesmus obliquus. Biochem. J. 1996, 316, 73–80. [Google Scholar] [CrossRef]
Kasahara, H.; Hanada, A.; Kuzuyama, T.; Takagi, M.; Kamiya, Y.; Yamaguchi, S. Contribution of the mevalonate and methylerythritol phosphate pathways to the biosynthesis of gibberellins in Arabidopsis. J. Biol. Chem. 2002, 277, 45188–45194. [Google Scholar] [CrossRef]
Cutler, A.J.; Krochko, J.E. Formation and breakdown of ABA. Trends Plant Sci. 1999, 4, 472–478. [Google Scholar] [CrossRef]
Hartung, W. The evolution of abscisic acid (ABA) and ABA function in lower plants fungi and lichen. Funct. Plant Biol. 2010, 37, 806–812. [Google Scholar] [CrossRef]
Seo, M.; Koshiba, T. Complex regulation of ABA biosynthesis in plants. Trends Plant Sci. 2002, 7, 41–48. [Google Scholar] [CrossRef]
Baroli, I.; Niyogi, K.K. Molecular genetics of xanthophyll–dependent photoprotection in green algae and plants. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2000, 355, 1385–1394. [Google Scholar] [CrossRef] [PubMed]
Watanabe, T.; Kondo, N. Ethylene evolution in marine algae and a proteinaceous inhibitor of ethylene biosynthesis from red alga. Plant Cell Physiol. 1976, 17, 1159–1166. [Google Scholar] [CrossRef]
Plettner, I.N.A.; Steinke, M.; Malin, G. Ethene (ethylene) production in the marine macroalga Ulva (Enteromorpha) intestinalis L. (Chlorophyta, Ulvophyceae): Effect of light-stress and co-production with dimethyl sulphide. Plant Cell Environ. 2005, 28, 1136–1145. [Google Scholar] [CrossRef]
Maillard, P.; Thepenier, C.; Gudin, C. Determination of an ethylene biosynthesis pathway in the unicellular green alga, Haematococcus pluvialis. Relationship between growth and ethylene production. J. Appl. Phycol. 1993, 5, 93–98. [Google Scholar] [CrossRef]
Booker, M.A.; DeLong, A. Producing the ethylene signal: Regulation and diversification of ethylene biosynthetic enzymes. Plant Physiol. 2015, 69, 42–50. [Google Scholar] [CrossRef] [PubMed]
Cembella, A.D. Chemical ecology of eukaryotic microalgae in marine ecosystems. Phycologia 2003, 42, 420–447. [Google Scholar] [CrossRef]
Frenkel, J.; Vyverman, W.; Pohnert, G. Pheromone signaling during sexual reproduction in algae. Plant J. 2014, 79, 632–644. [Google Scholar] [CrossRef] [PubMed]
Stratmann, K.; Boland, W.; Müller, D.G. Pheromones of marine brown algae; a new branch of the eicosanoid metabolism. Angew. Chem. Int. Ed. Engl. 1992, 31, 1246–1248. [Google Scholar] [CrossRef]
Boland, W.; Jaenicke, L.; Muller, D.G.; Gassmann, G. Giffordene, 2Z, 4Z, 6E, 8Z-undecatetraene, is the odoriferous principle of the marine brown alga Giffordia mitchellae. Experientia 1987, 43, 466–467. [Google Scholar] [CrossRef]
Muller, D.G.; Schmid, C.E. Qualitative and quantitative determination of pheromone secretion in female gametes of Ectocarpus siliculosus (Phaeophyceae). Biol. Chem. Hoppe-Seyler 1988, 369, 647–653. [Google Scholar] [CrossRef]
Stratmann, K.; Boland, W.; Müller, D.G. Biosynthesis of pheromones in female gametes of marine brown algae (Phaeophyceae). Tetrahedron 1993, 49, 3755–3766. [Google Scholar] [CrossRef]
Yannai, S. Dictionary of Food Compounds with CD-ROM: Additives, Flavors, and Ingredients, 1st ed.; Chapman & Hall/CRC: Boca Raton, FL, USA, 2004; p. 1784. [Google Scholar]
Rui, F.; Boland, W. Algal pheromone biosynthesis: Stereochemical analysis and mechanistic implications in gametes of Ectocarpus siliculosus. J. Org. Chem. 2010, 75, 3958–3964. [Google Scholar] [CrossRef] [PubMed]
Pohnert, G.; Boland, W. The oxylipin chemistry of attraction and defense in brown algae and diatoms. Nat. Prod. Rep. 2002, 19, 108–122. [Google Scholar] [CrossRef] [PubMed]
Boland, W. The chemistry of gamete attraction: Chemical structures, biosynthesis, and (a) biotic degradation of algal pheromones. Proc. Natl. Acad. Sci. USA 1995, 92, 37–43. [Google Scholar] [CrossRef] [PubMed]
Pohnert, G.; Boland, W. Biosynthesis of the algal pheromone hormosirene by the fresh-water diatom Gomphonema parvulum (Bacillariophyceae). Tetrahedron 1996, 52, 10073–10082. [Google Scholar] [CrossRef]
Kajiwara, T.; Kodama, K.; Hatanaka, A.; Matsui, K. Volatile Compounds from Japanese Marine Brown Algae. In Bioactive Volatile Compounds from Plants; Teranishi, R., Buttery, R.G., Sugisawa, H., Eds.; American Chemical Society: Washington, DC, USA, 1993; Volume 525, pp. 103–120. [Google Scholar] [CrossRef]
Moore, R.E. Volatile compounds from marine algae. Acc. Chem. Res. 1977, 10, 40–47. [Google Scholar] [CrossRef]
Gruen, H.E. Auxins and fungi. Annu. Rev. Plant Physiol. 1959, 10, 405–440. [Google Scholar] [CrossRef]
Spaepen, S.; Vanderleyden, J.; Remans, R. Indole-3-acetic acid in microbial and microorganism-plant signaling. FEMS Microbiol. Rev. 2007, 31, 425–448. [Google Scholar] [CrossRef]
Lau, S.; Shao, N.; Bock, R.; Jürgens, G.; De Smet, I. Auxin signaling in algal lineages: Fact or myth? Trends Plant Sci. 2009, 14, 182–188. [Google Scholar] [CrossRef]
Tarakhovskaya, E.R.; Maslov, Y.I.; Shishova, M.F. Phytohormones in algae. Russ. J. Plant Physiol. 2007, 54, 163–170. [Google Scholar] [CrossRef]
Yokoya, N.S.; Handro, W. Effects of auxins and cytokinins on tissue culture of Grateloupia dichotoma (Gigartinales, Rhodophyta). Hydrobiologia 1996, 326, 393–400. [Google Scholar] [CrossRef]
Polevoi, V.V.; Tarakhovskaya, E.R.; Maslov, Y.I.; Polevoi, A.V. Role of auxin in induction of polarity in Fucus vesiculosus zygotes. Russ. J. Dev. Biol. 2003, 34, 360–364. [Google Scholar] [CrossRef]
Fries, L. Growth regulating effects of phenylacetic acid and p-hydroxyphenylacetic acid on Fucus spiralis L. (Phaeophyceae, Fucales) in axenic culture. Phycologia 1977, 16, 451–455. [Google Scholar] [CrossRef]
Fries, L.; Iwasaki, H. p-Hydroxyphenylacetic acid and other phenolic compounds as growth stimulators of the red alga Porphyra tenera. Plant Sci. Lett. 1976, 6, 299–307. [Google Scholar] [CrossRef]
Buggeln, R.G. Morphogenesis and Growth Regulators. In The Biology of Seaweeds; Wiley-Blackwell: Oxford, UK, 1981; pp. 627–660. [Google Scholar]
Jacobs, W.P.; Falkenstein, K.; Hamilton, R.H. Nature and amount of auxin in algae: IAA from extracts of Caulerpa paspaloides (Siphonales). Plant Physiol. 1985, 78, 844–848. [Google Scholar] [CrossRef] [PubMed]
Chen, X.; Tang, Y.; Sun, X.; Zhang, X.; Xu, N. Comparative transcriptome analysis reveals the promoting effects of IAA on biomass production and branching of Gracilariopsis lemaneiformis. Aquaculture 2022, 548, 737678. [Google Scholar] [CrossRef]
Mooney, P.A.; Van Staden, J. Seasonal changes in the levels of endogenous cytokinins in Sargassum heterophyllum (Phaeophyceae). Bot. Mar. 1984, 27, 437–442. [Google Scholar] [CrossRef]
De Nys, R.; Jameson, P.E.; Chin, N.; Brown, M.T.; Sanderson, K.J. The cytokinins as endogenous growth regulators in Macrocystis pyrifera (L.) C. Ag. (Phaeophyceae). Bot. Mar. 1990, 33, 467–475. [Google Scholar] [CrossRef]
Radley, M. Gibberellin-like Substances in Plants: Leguminous Root Nodules. Nature 1961, 191, 684–685. [Google Scholar] [CrossRef]
Jacobs, W.P. A search for some angiosperm hormones and their metabolites in Caulerpa paspaloides (Chlorophyta). J. Phycol. 1993, 29, 595–600. [Google Scholar] [CrossRef]
Taylor, I.B.; Burbidge, A.; Thompson, A.J. Control of abscisic acid synthesis. J. Exp. Bot. 2000, 51, 1563–1574. [Google Scholar] [CrossRef] [PubMed]
Hirsch, R.; Hartung, W.; Gimmler, H. Abscisic acid content of algae under stress. Bot. Acta 1989, 102, 326–334. [Google Scholar] [CrossRef]
Nimura, K.; Mizuta, H. Inducible effects of abscisic acid on sporophyte discs from Laminaria japonica Areschoug (Laminariales, Phaeophyceae). J. Appl. Phycol. 2002, 14, 159–163. [Google Scholar] [CrossRef]
Uji, T.; Matsuda, R.; Takechi, K.; Takano, H.; Mizuta, H.; Takio, S. Ethylene regulation of sexual reproduction in the marine red alga Pyropia yezoensis (Rhodophyta). J. Appl. Phycol. 2016, 28, 3501–3509. [Google Scholar] [CrossRef]
Uji, T.; Mizuta, H. The role of plant hormones on the reproductive success of red and brown algae. Front. Plant Sci. 2022, 13, 1019334. [Google Scholar] [CrossRef] [PubMed]
Van de Poel, B.; Cooper, E.D.; Van Der Straeten, D.; Chang, C.; Delwiche, C.F. Transcriptome profiling of the green alga Spirogyra pratensis (Charophyta) suggests an ancestral role for ethylene in cell wall metabolism, photosynthesis, and abiotic stress responses. Plant Physiol. 2016, 172, 533–545. [Google Scholar] [CrossRef] [PubMed]
Bajguz, A.; Czerpak, R. Physiological and biochemical role of brassinosteroids and their structure-activity relationship in the green alga Chlorella vulgaris Beijerinck (Chlorophyceae). J. Plant Growth Regul. 1998, 17, 131–139. [Google Scholar] [CrossRef]
Bajguz, A. Effect of brassinosteroids on nucleic acids and protein content in cultured cells of Chlorella vulgaris. Plant Physiol. Biochem. 2000, 38, 209–215. [Google Scholar] [CrossRef]
Liu, J.; Qiu, W.; Xia, D. Brassinosteroid improves lipid productivity and stress tolerance of Chlorella cells induced by high temperature. J. Appl. Phycol. 2018, 30, 253–260. [Google Scholar] [CrossRef]
Bajguz, A. Brassinosteroid enhanced the level of abscisic acid in Chlorella vulgaris subjected to short-term heat stress. J. Plant Physiol. 2009, 166, 882–886. [Google Scholar] [CrossRef]
Kazan, K.; Lyons, R. Intervention of phytohormone pathways by pathogen effectors. Plant Cell 2014, 26, 2285–2309. [Google Scholar] [CrossRef] [PubMed]
Feys, B.J.; Parker, J.E. Interplay of signaling pathways in plant disease resistance. Trends Genet. 2000, 16, 449–455. [Google Scholar] [CrossRef] [PubMed]
Wiesemeier, T.; Jahn, K.; Pohnert, G. No evidence for the induction of brown algal chemical defense by the phytohormones jasmonic acid and methyl jasmonate. J. Chem. Ecol. 2008, 34, 1523–1531. [Google Scholar] [CrossRef] [PubMed]
Tabor, C.W.; Tabor, H. Polyamines. Annu. Rev. Biochem. 1984, 53, 749–790. [Google Scholar] [CrossRef] [PubMed]
Xu, B.; Bo, Y.; Sun, X.; Wang, H.; Guo, H.; Zhou, C.; Ruan, R.; Yan, X.; Cheng, P. Review of the effect of polyamines in microalgae when ingested by shellfish. Algal Res. 2021, 58, 102409. [Google Scholar] [CrossRef]
Lefevere, H.; Bauters, L.; Gheysen, G. Salicylic acid biosynthesis in plants. Front. Plant Sci. 2020, 11, 338. [Google Scholar] [CrossRef] [PubMed]
Faizan, M.; Faraz, A.; Sami, F.; Siddiqui, H.; Yusuf, M.; Gruszka, D.; Hayat, S. Role of strigolactones: Signalling and crosstalk with other phytohormones. Open Life Sci. 2020, 15, 217–228. [Google Scholar] [CrossRef] [PubMed]
Bouwmeester, H.J.; Roux, C.; Lopez-Raez, J.A.; Becard, G. Rhizosphere communication of plants, parasitic plants and AM fungi. Trends Plant Sci. 2007, 12, 224–230. [Google Scholar] [CrossRef]
Stirk, W.A.; Van Staden, J. Plant growth regulators in seaweeds: Occurrence, regulation and functions. Adv. Bot. Res. 2014, 71, 125–159. [Google Scholar] [CrossRef]
Arioli, T.; Mattner, S.W.; Winberg, P.C. Applications of seaweed extracts in Australian agriculture: Past, present and future. J. Appl. Phycol. 2015, 27, 2007–2015. [Google Scholar] [CrossRef]
Waaland, S.D.; Watson, B.A. Isolation of a cell-fusion hormone from Griffithsia pacifica Kylin, a red alga. Planta 1980, 149, 493–497. [Google Scholar] [CrossRef]
Wichard, T.; Oertel, W. Gametogenesis and gamete release of Ulva mutabilis and Ulva lactuca (Chlorophyta): Regulatory effects and chemical characterization of the “swarming inhibitor”. J. Phycol. 2010, 46, 248–259. [Google Scholar] [CrossRef]
Kamiya, Y. Plant hormones: Versatile regulators of plant growth and development. Annu. Rev. Plant Biol. 2010, 60. [Google Scholar] [CrossRef]
Jaillais, Y.; Chory, J. Unraveling the paradoxes of plant hormone signaling integration. Nat. Struct. Mol. Biol. 2010, 17, 642–645. [Google Scholar] [CrossRef] [PubMed]
Friml, J.; Jones, A.R. Endoplasmic reticulum: The rising compartment in auxin biology. Plant Physiol. 2010, 154, 458–462. [Google Scholar] [CrossRef] [PubMed]
Effendi, Y.; Scherer, G.F. AUXIN BINDING-PROTEIN1 (ABP1), a receptor to regulate auxin transport and early auxin genes in an interlocking system with PIN proteins and the receptor TIR1. Plant Signal. Behav. 2011, 6, 1101–1103. [Google Scholar] [CrossRef] [PubMed]
Gray, W.M.; Kepinski, S.; Rouse, D.; Leyser, O.; Estelle, M. Auxin regulates SCFTIR1-dependent degradation of AUX/IAA proteins. Nature 2001, 414, 271–276. [Google Scholar] [CrossRef] [PubMed]
Lu, Y.; Xu, J. Phytohormones in microalgae: A new opportunity for microalgal biotechnology? Trends Plant Sci. 2015, 20, 273–282. [Google Scholar] [CrossRef] [PubMed]
Dharmasiri, S.; Jayaweera, T.; Dharmasiri, N. Plant hormone signalling: Current perspectives on perception and mechanisms of action. Ceylon J. Sci. Biol. Sci. 2013, 42, 1–17. [Google Scholar] [CrossRef]
Lu, Y.; Tarkowská, D.; Turečková, V.; Luo, T.; Xin, Y.; Li, J.; Wang, Q.; Jiao, N.; Strnad, M.; Xu, J. Antagonistic roles of abscisic acid and cytokinin during response to nitrogen depletion in oleaginous microalga Nannochloropsis oceanica expand the evolutionary breadth of phytohormone function. Plant J. 2014, 80, 52–68. [Google Scholar] [CrossRef]
Schwechheimer, C.; Willige, B.C. Shedding light on gibberellic acid signalling. Curr. Opin. Plant Biol. 2009, 12, 57–62. [Google Scholar] [CrossRef] [PubMed]
Leivar, P.; Monte, E.; Oka, Y.; Liu, T.; Carle, C.; Castillon, A.; Huq, E.; Quail, P.H. Multiple phytochrome-interacting bHLH transcription factors repress premature seedling photomorphogenesis in darkness. Curr. Biol. 2008, 18, 1815–1823. [Google Scholar] [CrossRef]
Vandenbussche, F.; Fierro, A.C.; Wiedemann, G.; Reski, R.; van der Straeten, D. Evolutionary conservation of plant gibberellin signalling pathway components. BMC Plant Biol. 2007, 7, 65. [Google Scholar] [CrossRef] [PubMed]
Park, S.Y.; Fung, P.; Nishimura, N.; Jensen, D.R.; Fujii, H.; Zhao, Y.; Lumba, S.; Santiago, J.; Rodrigues, A.; Chow, T.F.; et al. Abscisic acid inhibits type 2C protein phosphatases via the PYR/PYL family of START proteins. Science 2009, 324, 1068–1071. [Google Scholar] [CrossRef] [PubMed]
Guo, J.; Yang, X.; Weston, D.J.; Chen, J.G. Abscisic acid receptors: Past, present and future. J. Integr. Plant Biol. 2011, 53, 469–479. [Google Scholar] [CrossRef] [PubMed]
Hauser, F.; Waadt, R.; Schroeder, J.I. Evolution of abscisic acid synthesis and signaling mechanisms. Curr. Biol. 2011, 21, R346–R355. [Google Scholar] [CrossRef] [PubMed]
Hanada, K.; Hase, T.; Toyoda, T.; Shinozaki, K.; Okamoto, M. Origin and evolution of genes related to ABA metabolism and its signaling pathways. J. Plant Res. 2011, 124, 455–465. [Google Scholar] [CrossRef] [PubMed]
Yoo, S.D.; Cho, Y.; Sheen, J. Emerging connections in the ethylene signaling network. Trends Plant Sci. 2009, 14, 270–279. [Google Scholar] [CrossRef]
Bleecker, A.B. Ethylene perception and signaling: An evolutionary perspective. Trends Plant Sci. 1999, 4, 269–274. [Google Scholar] [CrossRef]
Mori, I.C.; Ikeda, Y.; Matsuura, T.; Hirayama, T.; Mikami, K. Phytohormones in red seaweeds: A technical review of methods for analysis and a consideration of genomic data. Bot. Mar. 2017, 60, 153–170. [Google Scholar] [CrossRef]
Moon, J.S.; Kim, G.H. Somatic cell fusion in a red alga Griffithsia monilis is mediated by two different signalling molecules. Phycologia 2017, 56, 130. [Google Scholar]
Starr, R.C.; Marner, F.J.; Jaenicke, L. Chemoattraction of male gametes by a pheromone produced by female gametes of Chlamydomonas. Proc. Natl. Acad. Sci. USA 1995, 92, 641–645. [Google Scholar] [CrossRef] [PubMed]
Adair, W.S. Characterization of Chlamydomonas sexual agglutinins. J. Cell Sci. 1985, 1985 (Suppl. S2), 233–260. [Google Scholar] [CrossRef] [PubMed]
Mori, K.; Takanashi, S.I. Synthesis of lurlene, the sex pheromone of the green flagellate Chlamydomonas allensworthii. Tetrahedron Lett. 1996, 37, 1821–1824. [Google Scholar] [CrossRef]
Kirk, D.L.; Kirk, M.M. Heat shock elicits production of sexual inducer in Volvox. Science 1986, 231, 51–54. [Google Scholar] [CrossRef] [PubMed]
Godl, K.; Hallmann, A.; Rappel, A.; Sumper, M. Pherophorins: A family of extracellular matrix glycoproteins from Volvox structurally related to the sex-inducing pheromone. Planta 1995, 196, 781–787. [Google Scholar] [CrossRef] [PubMed]
Sumper, M.; Berg, E.; Wenzl, S.; Godl, K. How a sex pheromone might act at a concentration below 10–16 M. EMBO J. 1993, 12, 831–836. [Google Scholar] [CrossRef] [PubMed]
Amon, P.; Haas, E.; Sumper, M. The sex-inducing pheromone and wounding trigger the same set of genes in the multicellular green alga Volvox. Plant Cell 1998, 10, 781–789. [Google Scholar] [CrossRef]
Schmid, C.E. Cell-cell-recognition during fertilization in Ectocarpus siliculosus (Phaeophyceae). Hydrobiologia 1993, 260, 437–443. [Google Scholar] [CrossRef]
Maier, I.; Muller, D.G. Antheridium fine structure and spermatozoid release in Laminaria digitata (Phaeophyceae). Phycologia 1982, 21, 1–8. [Google Scholar] [CrossRef]
Maier, I.; Muller, D.G.; Schmid, C.; Boland, W.; Jaenicke, L. Pheromone receptor specificity and threshold concentrations for spermatozoid release in Laminaria digitata. Nat. Sci. 1988, 75, 260–263. [Google Scholar] [CrossRef]
Maier, I.; Muller, D.G. Chemotaxis in Laminaria digitata (Phaeophyceae) I. Analysis of spermatozoid movement. J. Exp. Bot. 1990, 41, 869–876. [Google Scholar] [CrossRef]
Maier, I.; Wenden, A.; Clayton, M.N. The movement of Hormosira banksii (Fucales, Phaeophyta) spermatozoids in response to sexual pheromone. J. Exp. Bot. 1992, 43, 1651–1657. [Google Scholar] [CrossRef]
Wirth, D.; Boland, W. Structure and Synthesis of (±)-Caudoxirene, a New Spermatozoid-Releasing and-Attracting Pheromone from the Marine Brown Alga Perithalia caudata (Phaeophyceae, Sporochnales). Helv. Chim. Acta 1990, 73, 916–921. [Google Scholar] [CrossRef]
Boland, W.; Hoever, F.P.; Kruger, B.W. Application of Molecular Modelling Techniques to Pheromones of the Marine Brown Algae Cutleria multifida and Ectocarpus siliculosus (Phaeophyceae). Metalloproteins as Chemoreceptors? J. Nat. Sci. 1989, 44, 29–837. [Google Scholar] [CrossRef]
Kadam, S.U.; Tiwari, B.K.; O’Donnell, C.P. Application of novel extraction technologies for bioactives from marine algae. J. Agric. Food Chem. 2013, 61, 4667–4675. [Google Scholar] [CrossRef] [PubMed]
Herrero, M.; del Pilar Sánchez-Camargo, A.; Cifuentes, A.; Ibáñez, E. Plants, seaweeds, microalgae and food by-products as natural sources of functional ingredients obtained using pressurized liquid extraction and supercritical fluid extraction. TrAC Trends Anal. Chem. 2015, 71, 26–38. [Google Scholar] [CrossRef]
Górka, B.; Wieczorek, P.P. Simultaneous determination of nine phytohormones in seaweed and algae extracts by HPLC-PDA. J. Chromatogr. B 2017, 1057, 32–39. [Google Scholar] [CrossRef]
Lu, Q.; Chen, L.; Lu, M.; Chen, G.; Zhang, L. Extraction and analysis of auxins in plants using dispersive liquid−liquid microextraction followed by high-performance liquid chromatography with fluorescence detection. J. Agric. Food Chem. 2010, 58, 2763–2770. [Google Scholar] [CrossRef]
Gupta, V.; Kumar, M.; Brahmbhatt, H.; Reddy, C.R.K.; Seth, A.; Jha, B. Simultaneous determination of different endogenetic plant growth regulators in common green seaweeds using dispersive liquid–liquid microextraction method. Plant Physiol. Biochem. 2011, 49, 1259–1263. [Google Scholar] [CrossRef]
Mikami, K.; Mori, I.C.; Matsuura, T.; Ikeda, Y.; Kojima, M.; Sakakibara, H.; Hirayama, T. Comprehensive quantification and genome survey reveal the presence of novel phytohormone action modes in red seaweeds. J. Appl. Phycol. 2016, 28, 2539–2548. [Google Scholar] [CrossRef]
Benítez García, I.; Dueñas Ledezma, A.K.; Martínez Montaño, E.; Salazar Leyva, J.A.; Carrera, E.; Osuna Ruiz, I. Identification and quantification of plant growth regulators and antioxidant compounds in aqueous extracts of Padina durvillaei and Ulva lactuca. Agronomy 2020, 10, 866. [Google Scholar] [CrossRef]
Stirk, W.A.; Bálint, P.; Tarkowská, D.; Novák, O.; Maróti, G.; Ljung, K.; Turečková, V.; Strnad, M.; Ördög, V.; van Staden, J. Effect of light on growth and endogenous hormones in Chlorella minutissima (Trebouxiophyceae). Plant Physiol. Biochem. 2014, 79, 66–76. [Google Scholar] [CrossRef] [PubMed]
Stirk, W.A.; Bálint, P.; Tarkowská, D.; Novák, O.; Strnad, M.; Ördög, V.; van Staden, J. Hormone profiles in microalgae: Gibberellins and brassinosteroids. Plant Physiol. Biochem. 2013, 70, 348–353. [Google Scholar] [CrossRef] [PubMed]
Stirk, W.A.; Ördög, V.; Novák, O.; Rolčík, J.; Strnad, M.; Bálint, P.; van Staden, J. Auxin and cytokinin relationships in 24 microalgal strains. J. Phycol. 2013, 49, 459–467. [Google Scholar] [CrossRef] [PubMed]
Grob, K.; Zürcher, F. Stripping of trace organic substances from water: Equipment and procedure. J. Chromatogr. A 1976, 17, 285–294. [Google Scholar] [CrossRef]
Boland, W.; Ney, P.; Jaenicke, L.; Gassmann, G. A “Closed-Loop-Stripping” Technique as a Versatile Tool for Metabolic Studies of Volatiles. In Analysis of Volatiles: Method, Application; Schreiber, P., Ed.; Walter de Gruyter GmbH: Berlin, Germany, 1984; pp. 371–380. [Google Scholar]
Maier, I.; Müller, D.G.; Gassmann, G.; Boland, W.; Jaenicke, L. Sexual pheromones and related egg secretions in Laminariales (Phaeophyta). J. Nat. Sci. 1987, 42, 948–954. [Google Scholar] [CrossRef]
Maier, I.; Clayton, M.N. Quantitative evaluation of erotactin secretion in eggs of Hormosira banksii (Fucales, Phaeophyceae). Bot. Acta 1993, 106, 344–349. [Google Scholar] [CrossRef]
Akakabe, Y.; Kajiwara, T. Bioactive volatile compounds from marine algae: Feeding attractants. In Nineteenth International Seaweed Symposium: Proceedings of the 19th International Seaweed Symposium, Kobe, Japan, 26–31 March 2007; Borowitzka, M.A., Critchley, A.T., Kraan, S., Peters, A., Sjøtun, K., Notoya, M., Eds.; Springer: Dordrecht, The Netherlands, 2009; pp. 211–214. [Google Scholar] [CrossRef]
Redshaw, E.S.; Hougen, F.W.; Baker, R.J. Distillation technique for isolation of volatile materials for gas chromatographic analysis and its application to coriander seed (Coriandrum sativum). J. Agric. Food Chem. 1971, 19, 1264–1266. [Google Scholar] [CrossRef]
Derenbach, J.B.; Pesando, D. Investigations into a small fraction of volatile hydrocarbons: III. Two diatom cultures produce ectocarpene, a pheromone of brown algae. Mar. Chem. 1986, 19, 337–341. [Google Scholar] [CrossRef]
Weber, R.J.; Selander, E.; Sommer, U.; Viant, M.R. A stable-isotope mass spectrometry-based metabolic footprinting approach to analyze exudates from phytoplankton. Mar. Drugs 2013, 11, 4158–4175. [Google Scholar] [CrossRef] [PubMed]
Hurtado, A.Q.; Neish, I.; Khan, M.; Norrie, J.; Pereira, L.; Michalak, I.; Shukla, P.S.; Critchley, A.T. Extracts of seaweeds used as biostimulatns on land and sea crops—An efficacious, phyconomic, circular blue economy: With special reference to Ascophyllum (brown) and Kappaphycus (red) seaweeds. In Biostimulants for Crops from Seed Germination to Plant Development: A Practical Approach; Gupta, S., Van Staden, J., Eds.; Academic Press: Cambridge, MA, USA, 2021; pp. 263–288. [Google Scholar] [CrossRef]
Sati, H.; Chokshi, K.; Soundarya, R.; Ghosh, A.; Mishra, S. Seaweed-based biostimulant improves photosynthesis and effectively enhances growth and biofuel potential of a green microalga Chlorella variabilis. Aquac. Int. 2021, 29, 963–975. [Google Scholar] [CrossRef]
Pilar, G.J.; Olegario, B.R.; Rafael, R.R. Occurrence of jasmonates during cystocarp development in the red alga Grateloupia imbricata. J. Phycol. 2016, 52, 1085–1093. [Google Scholar] [CrossRef] [PubMed]
García-Jiménez, P.; Robaina, R.R. Effects of ethylene on tetrasporogenesis in Pterocladiella capillacea (Rhodophyta). J. Phycol. 2012, 48, 710–715. [Google Scholar] [CrossRef] [PubMed]
Uji, T.; Endo, H.; Mizuta, H. Sexual reproduction via a 1-aminocyclopropane-1-carboxylic acid-dependent pathway through redox modulation in the marine red alga Pyropia yezoensis (Rhodophyta). Front. Plant Sci. 2020, 11, 60. [Google Scholar] [CrossRef] [PubMed]
Guajardo, E.; Correa, J.A.; Contreras-Porcia, L. Role of abscisic acid (ABA) in activating antioxidant tolerance responses to desiccation stress in intertidal seaweed species. Planta 2016, 243, 767–781. [Google Scholar] [CrossRef] [PubMed]
Yokoya, N.S.; West, J.A.; Luchi, A.E. Effects of plant growth regulators on callus formation, growth and regeneration in axenic tissue cultures of Gracilaria tenuistipitata and Gracilaria perplexa (Gracilariales, Rhodophyta). Phycol. Res. 2004, 52, 244–254. [Google Scholar] [CrossRef]
Yeong, H.Y.; Phang, S.M.; Reddy, C.R.K.; Khalid, N. Production of clonal planting materials from Gracilaria changii and Kappaphycus alvarezii through tissue culture and culture of G. changii explants in airlift photobioreactors. J. Appl. Phycol. 2014, 26, 729–746. [Google Scholar] [CrossRef]
Kazi, M.A.; Singh, A.; Grewal, M.; Baraiya, M.; Goswami, S.; Rathore, M.S.; Jaiswar, S.; Mantri, V.A. Comparative evaluation of bio-effectors on survival and regeneration in Gracilaria dura (Rhodophyta). J. Appl. Phycol. 2022, 34, 3127–3139. [Google Scholar] [CrossRef]
Hurtado, A.Q.; Cheney, D.P. Propagule production of Eucheuma denticulatum (Burman) Collins et Harvey by tissue culture. Bot. Mar. 2003, 46, 338–341. [Google Scholar] [CrossRef]
Aguirre-Lipperheide, M.; Estrada-Rodriyuez, F.J.; Evans, L.V. Facts, problems, and needs in seaweed tissue culture: An appraisal. J. Phycol. 1995, 31, 677–688. [Google Scholar] [CrossRef]
Patwary, M.U.; Van der Meer, J.P. Construction of backcrossed Gelidium male-sterile and male-fertile lines and their growth comparison. J. Appl. Phycol. 1997, 8, 483–486. [Google Scholar] [CrossRef]
Luo, L.; Zuo, X.; Guo, L.; Pang, G.; Ma, Z.; Wu, M.; Chen, B. Effects of exogenous hormones on the regeneration of juveniles from Sargassum fusiforme holdfasts. Front. Mar. Sci. 2023, 9, 1072391. [Google Scholar] [CrossRef]
Hurtado, A.Q.; Biter, A.B. Plantlet regeneration of Kappaphycus alvarezii var. adik-adik by tissue culture. J. Appl. Phycol. 2007, 19, 783–786. [Google Scholar] [CrossRef]
Hurtado, A.Q.; Neish, I.C.; Critchley, A.T. Developments in production technology of Kappaphycus in the Philippines: More than four decades of farming. J. Appl. Phycol. 2015, 27, 1945–1961. [Google Scholar] [CrossRef]
Souza, J.M.; Castro, J.Z.; Critchley, A.T.; Yokoya, N.S. Physiological responses of the red algae Gracilaria caudata (Gracilariales) and Laurencia catarinensis (Ceramiales) following treatment with a commercial extract of the brown alga Ascophyllum nodosum (AMPEP). J. Appl. Phycol. 2019, 31, 1883–1888. [Google Scholar] [CrossRef]
Dawange, P.; Jaiswar, S. Effects of Ascophyllum marine plant extract powder (AMPEP) on tissue growth, proximate, phenolic contents, and free radical scavenging activities in endemic red seaweed Gracilaria corticata var. cylindrica from India. J. Appl. Phycol. 2020, 32, 4127–4135. [Google Scholar] [CrossRef]
Panda, D.; Pramanik, K.; Nayak, B.R. Use of sea weed extracts as plant growth regulators for sustainable agriculture. Int. J. Bioresour. Stress Manag. 2012, 3, 404–411. [Google Scholar]
Kozlova, T.A.; Hardy, B.P.; Krishna, P.; Levin, D.B. Effect of phytohormones on growth and accumulation of pigments and fatty acids in the microalgae Scenedesmus quadricauda. Algal Res. 2017, 27, 325–334. [Google Scholar] [CrossRef]
Hunt, R.W.; Chinnasamy, S.; Bhatnagar, A.; Das, K.C. Effect of biochemical stimulants on biomass productivity and metabolite content of the microalga, Chlorella sorokiniana. Appl. Biochem. Biotechnol. 2010, 162, 2400–2414. [Google Scholar] [CrossRef]
Han, X.; Zeng, H.; Bartocci, P.; Fantozzi, F.; Yan, Y. Phytohormones and effects on growth and metabolites of microalgae: A review. Fermentation 2018, 4, 25. [Google Scholar] [CrossRef]
Motomura, T.; Sakai, Y. The occurrence of flagellated eggs in Laminaria angustata (Phaeophyta, Laminariales). J. Phycol. 1988, 24, 282–285. [Google Scholar] [CrossRef]
Brawley, S.H.; Johnson, L.E. Gametogenesis, gametes and zygotes: An ecological perspective on sexual reproduction in the algae. Br. Phycol. J. 1992, 27, 233–252. [Google Scholar] [CrossRef]
Moeys, S.; Frenkel, J.; Lembke, C.; Gillard, J.T.; Devos, V.; Van den Berge, K.; Bouillon, B.; Huysman, M.J.; De Decker, S.; Scharf, J.; et al. A sex-inducing pheromone triggers cell cycle arrest and mate attraction in the diatom Seminavis robusta. Sci. Rep. 2016, 6, 19252. [Google Scholar] [CrossRef] [PubMed]
Garcia-Jimenez, P.; Montero-Fernández, M.; Robaina, R.R. Analysis of ethylene-induced gene regulation during carposporogenesis in the red seaweed Grateloupia imbricata (Rhodophyta). J. Phycol. 2018, 54, 681–689. [Google Scholar] [CrossRef] [PubMed]
El Shoubaky, G.A.; Salem, E.A. Effect of abiotic stress on endogenous phytohormones profile in some seaweeds. IJPPR 2016, 8, 124–134. [Google Scholar]
Kothari, R.; Singh, H.M.; Azam, R.; Goria, K.; Bharti, A.; Singh, A.; Bajar, S.; Pathak, A.; Pandey, A.K.; Tyagi, V.V. Potential avenue of genetic engineered algal derived bioactive compounds: Influencing parameters, challenges and future prospects. Phytochem. Rev. 2023, 22, 935–968. [Google Scholar] [CrossRef]
Foo, E.; Plett, J.M.; Lopez-Raez, J.A.; Reid, D. The role of plant hormones in plant-microbe symbioses. Front. Plant Sci. 2019, 10, 1391. [Google Scholar] [CrossRef]
Singh, R.P.; Reddy, C.R.K. Seaweed-microbial interactions: Key functions of seaweed-associated bacteria. FEMS Microbiol. Ecol. 2014, 88, 213–230. [Google Scholar] [CrossRef]
Zhang, R.; Wang, B.; Ouyang, J.; Li, J.; Wang, Y. Arabidopsis indole synthase, a homolog of tryptophan synthase alpha, is an enzyme involved in the trp-independent indole-containing metabolite biosynthesis. J. Integr. Plant Biol. 2008, 50, 1070–1077. [Google Scholar] [CrossRef]
Ljung, K.; Hull, A.K.; Kowalczyk, M.; Marchant, A.; Celenza, J.; Cohen, J.D.; Sandberg, G. Biosynthesis, conjugation, catabolism and homeostasis of indole-3-acetic acid in Arabidopsis thaliana. Plant Mol. Biol. 2002, 49, 249–272. [Google Scholar] [CrossRef] [PubMed]
Bajguz, A.; Piotrowska-Niczyporuk, A. Synergistic effect of auxins and brassinosteroids on the growth and regulation of metabolite content in the green alga Chlorella vulgaris (Trebouxiophyceae). Plant Physiol. Biochem. 2013, 71, 290–297. [Google Scholar] [CrossRef] [PubMed]
Schaller, F.; Schaller, A.; Stintzi, A. Biosynthesis and metabolism of jasmonates. J. Plant Growth Regul. 2004, 23, 179–199. [Google Scholar] [CrossRef]
Vick, B.A.; Zimmerman, D.C. The biosynthesis of jasmonic acid: A physiological role for plant lipoxygenase. Biochem. Biophys. Res. Commun. 1983, 111, 470–477. [Google Scholar] [CrossRef] [PubMed]
Hamberg, M.; Hughes, M.A. Fatty acid allene oxides III Albumin-induced cyclization of 12, 13 (S)-epoxy-9 (Z), 11-octadecadienoic acid. Lipids 1988, 23, 469–475. [Google Scholar] [CrossRef]
Stumpe, M.; Göbel, C.; Faltin, B.; Beike, A.K.; Hause, B.; Himmelsbach, K.; Bode, J.; Kramell, R.; Wasternack, C.; Frank, W.; et al. The moss Physcomitrella patens contains cyclopentenones but no jasmonates: Mutations in allene oxide cyclase lead to reduced fertility and altered sporophyte morphology. New Phytol. 2010, 188, 740–749. [Google Scholar] [CrossRef] [PubMed]
Gundlach, H.; Zenk, M.H. Biological activity and biosynthesis of pentacyclic oxylipins: The linoleic acid pathway. Phytochemistry 1998, 47, 527–537. [Google Scholar] [CrossRef]
Collén, J.; Porcel, B.; Carré, W.; Ball, S.G.; Chaparro, C.; Tonon, T.; Barbeyron, T.; Michel, G.; Noel, B.; Valentin, K.; et al. Genome structure and metabolic features in the red seaweed Chondrus crispus shed light on evolution of the Archaeplastida. Proc. Natl. Acad. Sci. USA 2013, 110, 5247–5252. [Google Scholar] [CrossRef]
Hamberg, M.; Gerwick, W.H. Biosynthesis of vicinal dihydroxy fatty acids in the red alga Gracilariopsis lemaneiformis: Identification of a sodium-dependent 12-lipoxygenase and a hydroperoxide isomerase. Arch. Biochem. Biophys. 1993, 305, 115–122. [Google Scholar] [CrossRef]
Bagni, N.; Tassoni, A. Biosynthesis, oxidation and conjugation of aliphatic polyamines in higher plants. Amino Acids 2001, 20, 301–317. [Google Scholar] [CrossRef]
Liu, J.H.; Kitashiba, H.; Wang, J.; Ban, Y.; Moriguchi, T. Polyamines and their ability to provide environmental stress tolerance to plants. Plant Biotechnol. 2007, 24, 117–126. [Google Scholar] [CrossRef]
Kumar, A.; Taylor, M.; Altabella, T.; Tiburcio, A.F. Recent advances in polyamine research. Trends Plant Sci. 1997, 2, 124–130. [Google Scholar] [CrossRef]
Dixon, R.A.; Paiva, N.L. Stress-induced phenylpropanoid metabolism. Plant Cell 1995, 7, 1085. [Google Scholar] [CrossRef] [PubMed]
Yoneyama, K.; Xie, X.; Yoneyama, K.; Kisugi, T.; Nomura, T.; Nakatani, Y.; Akiyama, K.; McErlean, C.S. Which are the major players, canonical or non-canonical strigolactones? J. Exp. Bot. 2018, 69, 2231–2239. [Google Scholar] [CrossRef] [PubMed]
Chesterfield, R. A Synthetic Biology Toolbox for Examining and Engineering Strigolactone Biosynthesis. Ph.D. Thesis, The University of Queensland, Brisbane, Australia, 2020. [Google Scholar]
Sun, T.; Yuan, H.; Cao, H.; Yazdani, M.; Tadmor, Y.; Li, L. Carotenoid metabolism in plants: The role of plastids. Mol. Plant 2018, 11, 58–74. [Google Scholar] [CrossRef] [PubMed]
Alder, A.; Jamil, M.; Marzorati, M.; Bruno, M.; Vermathen, M.; Bigler, P.; Ghisla, S.; Bouwmeester, H.; Beyer, P.; Al-Babili, S. The path from β-carotene to carlactone, a strigolactone-like plant hormone. Science 2012, 335, 1348–1351. [Google Scholar] [CrossRef] [PubMed]
Kim, T.W.; Wang, Z.Y. Brassinosteroid signal transduction from receptor kinases to transcription factors. Annu. Rev. Plant Biol. 2010, 61, 681–704. [Google Scholar] [CrossRef]
de Saint Germain, A.; Ligerot, Y.; Dun, E.A.; Pillot, J.P.; Ross, J.J.; Beveridge, C.A.; Rameau, C. Strigolactones stimulate internode elongation independently of gibberellins. Plant Physiol. 2013, 163, 1012–1025. [Google Scholar] [CrossRef]
Wang, C.; Liu, Y.; Li, S.S.; Han, G.Z. Insights into the origin and evolution of the plant hormone signaling machinery. Plant Physiol. 2015, 167, 872–886. [Google Scholar] [CrossRef]
Fu, Z.Q.; Yan, S.; Saleh, A.; Wang, W.; Ruble, J.; Oka, N.; Mohan, R.; Spoel, S.H.; Tada, Y.; Zheng, N.; et al. NPR3 and NPR4 are receptors for the immune signal salicylic acid in plants. Nature 2012, 486, 228–232. [Google Scholar] [CrossRef]
Kim, G.H.; Fritz, L.A. Signal glycoprotein with α-d-mannosyl residues is involved in the wound-healing response of Antithamnion sparsum (ceramiales, rhodophyta). J. Phycol. 1993, 29, 85–90. [Google Scholar] [CrossRef]
Riad, N.; Reda Zahi, M.R.; Trovato, E.; Bouzidi, N.; Daghbouche, Y.; Utczás, M.; Mondello, L.; El Hattab, M. Chemical Screening and Antibacterial Activity of Essential Oil and Volatile Fraction of Dictyopteris polypodioides. Microchem. J. 2020, 152, 104415. [Google Scholar] [CrossRef]
Zhao, B.; Li, J. Regulation of brassinosteroid biosynthesis and inactivation. J. Integr. Plant Biol. 2012, 54, 746–759. [Google Scholar] [CrossRef] [PubMed]
Fujioka, S.; Yokota, T. Biosynthesis and metabolism of brassinosteroids. Annu. Rev. Plant Biol. 2003, 54, 137–164. [Google Scholar] [CrossRef] [PubMed]
Yan, Y.; Borrego, E.; Kolomiets, M.V. Jasmonate biosynthesis, perception and function in plant development and stress responses. In Lipid Metabolism; Baez, R.V., Ed.; InTech: London, UK, 2013; pp. 393–442. [Google Scholar]
León, J.; Sánchez-Serrano, J.J. Molecular biology of jasmonic acid biosynthesis in plants. Plant Physiol. Biochem. 1999, 37, 373–380. [Google Scholar] [CrossRef]
Kusano, T.; Berberich, T.; Tateda, C.; Takahashi, Y. Polyamines: Essential factors for growth and survival. Planta 2008, 228, 367–381. [Google Scholar] [CrossRef] [PubMed]
Bitrián, M.; Zarza, X.; Altabella, T.; Tiburcio, A.F.; Alcázar, R. Polyamines under abiotic stress: Metabolic crossroads and hormonal crosstalks in plants. Metabolites 2012, 2, 516–528. [Google Scholar] [CrossRef] [PubMed]
Kuznetsov, V.V.; Shevyakova, N.I. Polyamines and stress tolerance of plants. Plant Stress 2007, 1, 50–71. [Google Scholar]
Dempsey, D.M.A.; Vlot, A.C.; Wildermuth, M.C.; Klessig, D.F. Salicylic acid biosynthesis and metabolism. Arab. Book Am. Soc. Plant Biol. 2011, 9, e0156. [Google Scholar] [CrossRef]
Wani, K.I.; Chaudhary, S.; Zehra, A.; Naeem, M.; Aftab, T. Precise Role of Strigolactones and Its Crosstalk Mechanisms in Root Development. In Rhizobiology: Molecular Physiology of Plant Roots; Springer: Cham, Switzerland, 2021; pp. 253–270. [Google Scholar] [CrossRef]

Figure 1. Tryptophan-dependent biosynthesis of IAA (modified from [78,79]). Solid arrows: enzymes/genes responsible for the steps identified. Dashed arrows: proposed steps, but enzymes/genes have not been conclusively determined. Black color: pathway reported in algae. Blue color: pathway reported in higher plants.

Figure 2. Biosynthesis of cytokinin (modified from [78,84]).

Figure 3. Biosynthesis of gibberellin (modified from [78,84]). Black color: pathway reported in algae. Blue color: pathway reported in higher plants.

Figure 4. Biosynthesis of abscisic acid (modified from [78,84]).

Figure 5. Biosynthesis of ethylene (modified from [92,94]).

Figure 6. Biosynthesis of giffordene [98].

Figure 8. Biosynthesis of ectocarpene in brown algae from (9-hydroperoxyicosa-(5Z,7E,11Z,14Z,17Z)-pentaenoic acid (9-HPEPE) (modified from [104,105]). Black color: pathway found in algae. Blue color: pathway reported in higher plants.

Figure 9. Biosynthesis of dictyopterene [102].

Table 1. Role of hormones in algae.

Category	Hormone Name	Name of Alga	Role	Reference
Auxin	Phenylacetic acid (PAA)	Ulva compressa (as Enteromorpha compressa)	Induces the thallus growth	[24]
	Indole acetic acid (IAA)	Neoporphyra perforata (as Porphyra perforata)	Stimulates the growth and cell division	[25]
	Indole acetic acid (IAA)	Ulva lactuca	Induces the filamentous sporelings and promotes their growth	[26]
	Indole acetic acid (IAA)	Chara zeylanica	Antagonistic effect, inhibits gibberellic acid and its stimulatory effect on the growth	[27]
	Indole-3-acetic acid (IAA)	Caulerpa paspaloides	Enhanced initiation of leaf-like structures	[28]
	Indole acetic acid (IAA)	Fucus spiralis	Tissue differentiation	[29]
	Indole acetic acid (IAA)	Tretraselmis sps.	Tissue differentiation	[29]
Cytokinin	Isoprenoid and aromatic cytokinins	Cladophora capensis	Growth and morphogenesis	[30]
		Ulva sp.
		Caulerpa tongaensis (as Caulerpa filiformis)
		Halimeda cuneata
		Brassicophycus sisymbrioides (as Bifurcaria brassicaeformis)
		Ecklonia maxima
		Laminaria pallida
		Macrocystis pyrifera (as Macrocystis angustifolia)
		Splachnidium rugosum
		Dictyota sp.
		Sargassum incisifolium (as Sargassum heterophyllum)
		Pachymenia orbitosa (as Aeodes orbitosa)
		Gigartina bracteata (as Gigartina clathrata)
		Gigartina polycarpa
		Sarcothalia scutellata
		Hymenena venosa
		Hypnea spicifera
		Mazzaella capensis
		Nothogenia erinacea
		Plocamium corallorhiza
		Carradoeriella virgata
		Porphyra capensis
		Sarcothalia stiriata
		Gelidium vittatum (as Suhria vittata)
		Amphiroa bowerbankii
		Amphiroa ephedraea
		Arthrocardia sp.
		Cheilosporum sp.
		Corallina sp.
		Jania sp.
	Kinetin	Sphacelaria rigidula (as Sphacelaria furcigera)	Increases the length of lateral branches	[31]
	Kinetin	Kappaphycus alvarezii	Promotes cell division	[32]
	Zeatin	Kappaphycus alvarezii
		Sargassum tenerrimum
		Hydropuntia edulis (as Gracilaria edulis)
	Isopentenyladenine	Sargassum muticum	Early growth of the receptacles	[33]
	cis-zeatin riboside	Sargassum muticum	Early growth of the receptacles
	Isopentenyladenine	Neoporphyra perforata (as Porphyra perforata)	Helps in maturation of spermatangia and carpogonia
	cis-zeatin riboside	Neoporphyra perforata (as Porphyra perforata)	Helps in maturation of spermatangia and carpogonia
	Cytokinin	Fucus vesiculosus	Regeneration and morphogenesis	[34]
Gibberellic acid	Gibberellin (GA1/GA3)	Fucus spiralis	Tissue differentiation	[29]
	Gibberellin (GA4/GA7)	Tretraselmis sps.	Tissue differentiation	[29]
	Gibberellin (GA3)	Chara vulgaris	Promotes the number of antheridial filaments and spermatids	[35]
	Gibberellin (GA3)	Fucus vesiculosus	Increases adventitious branches	[34]
Abscisic acid	Abscisic acid (ABA)	Ulva lactuca/Ulva linza (as Ulva fasciata)	Releases in extreme conditions and inhibits growth	[36]
	Abscisic acid (ABA)	Dictyota humifusa	Releases in extreme conditions and inhibits growth	[36]
	Abscisic acid (ABA)	Phycocalidia acanthophora (as Porphyra acanthophora (red))	Releases in stress conditions and inhibits growth	[37]
		Phycocalidia acanthophora (as Porphyra acanthophora)
		Gelidium floridanum
		Crassiphycus birdiae (as Gracilaria birdiae)
		Gracilaria cervicornis
		Gracilariopsis tenuifrons
		Chondracanthus teedei
		Hypnea nigrescens
		Hypnea musciformis (brown)
		Hypnea musciformis (green)
		Dunaliella parva	Releases during salinity stress, causes reduction in growth, and in some cases promotes senescence	[38]
		Draparnaldia mutabilis
		Dunaliella acidophila	Reduction in growth and photosynthesis with increase in pH
Rhodomorphin	Rhodomorphin	Griffithsia pacifica	Repairs shoot cell and normal regeneration	[39]
		Anotrichium tenue
		Antithamnion kylinii
	Rhodomorphin (as glycoprotein)	Volvox carteri	Sexual hormone	[40]
Ethylene	1-aminocyclopropane-l-carboxylicacid (ACC) (Ethylene precursor)	Neoporphyra perforata (as Porphyra perforata)	Stimulates cell division and apical cap development	[25]
Ethylene	Ethylene	Acetabularia acetabulum (as Acetabularia mediterranea)	Influences the developmental pattern of cap	[41]
Brassinosteroids	Brassinolide (BR)	Ecklonia maxima	Improves stress response to various biotic and abiotic stresses	[42]
Brassinosteroids	Castasterone (CS)	Ecklonia maxima		[42]
Jasmonic acid	Jasmonate	Scenedesmus incrassulatus	Provides tolerance to the temperatures and infections stress	[43]
	Methyl jasmonate	Scenedesmus incrassulatus		[43]
	Jasmonic acid and its derivatives	Chondrus crispus	Induces defense reaction	[44]
Polyamines	Polyamines	Chlorella vulgaris	Enhances cell division, DNA replication, and autospore release	[45]
	Polyamines	Chlamydomonas reinhardtii	Enhances cell division	[46]
	Putrescine	Ulva lactuca (as Ulva fasciata)	Releases during hyposaline stress causes and decreases the chlorophyll and growth rate	[47]
	Spermidine	Ulva lactuca (as Ulva fasciata)		[47]
	Putrescine	Grateloupia doryphora	Releases during hyposaline shock and helps in physiological performance during acclimation by increasing photosynthetic rate	[48]
	Spermidine
	Spermine
	Polyamines	Grateloupia doryphora	Enhances cell division, elongation, and morphogenesis	[49,50]
	Putrescine	Grateloupia sp.	Induction of cystocarp, development, and release	[51]
	Spermidine
	Spermine
	Putrescine	Gracilaria cornea (as Crassiphycus corneus)	Promotes cystocarp maturation and liberation and develops cell masses	[52,53]
Salicylic acid	Salicylic acid	Saccharina japonica (as Laminaria japonica)	Imparts thermotolerance	[54]
Strigolactone	Strigolactone	Chara coralina	Stimulates rhizoid elongation	[23]

Table 2. Role of pheromones in algae.

Class	Name of Pheromone	Name of Alga	Role	Reference
Chlorophyceae	Sporulation inhibitor-1a (Glycoprotein)	Ulva compressa (as Ulva mutabilis)	Suppresses gametogenesis	[60]
	Swarming inhibitor	Ulva compressa (as Ulva mutabilis)	Inhibits gamete swarming
	Sporulation inhibitor-2 (Non-protein)	Ulva compressa (as Ulva mutabilis)	Suppresses gametogenesis
Phaeophyceae	Ectocarpene (S(+)l-cis-buten-l-yl-cyclohepta-2,5-diene)	Ectocarpus siliculosus	Releases female gamete and helps in attracting male gametes	[61]
		Sphacelaria rigidula	Acts as a chemoattractant	[62]
		Adenocystis longissima (as Adenocystis utricularis)	Acts as a chemoattractant	[63]
	Dictyotene (C11 metabolite)	Dictyopteris polypodioides (as Dictyopteris membranacea)	Helps in gamete attraction and acts as a deterrent to mesograzers	[64]
		Dictyota dichotoma
		Dictyota diemensis
		Sargassum filipendula
	C11 sulphur metabolite	Dictyopteris polypodioides (as Dictyopteris membranacea)
	Dictyotene (C11 metabolite)	Dictyota dimensis	Acts as a sperm attractant	[65]
	Diterpene alcohols	Dictyota dichotoma	Acts as a deterrent to herbivores	[66]
	C11 hydrocarbons	Dictyopteris delicatula	Acts as a deterrent to herbivores	[67]
	Thiopyranone (Thiopyran-4-one)	Dictyopteris polypodioides (as Dictyopteris membranacea)	Acts as a deterrent to herbivores	[68]
	Dithiepanone (Dithiepan-5-one)	Dictyopteris polypodioides (as Dictyopteris membranacea)	Acts as a deterrent to herbivores	[68]
	Multifidene	Halosiphon tomentosus (as Chorda tomentosa)	Acts as a chemoattractant	[69]
	Ectocarpene
	Dictyopterene
	Viridiene
	Fucoserratene (1,3-trans-5-cis-octatriene)	Fucus serratus	Acts as a chemoattractant	[70]
		Fucus vesiculosus		[71]
		Fucus spiralis		[71]
	Finavarrene	Ascophyllum nodosum	Acts as a chemoattractant	[71]
	Cystophorene	Cystophora siliquosa	Acts as a chemoattractant	[71]
	Hormosirene	Hormosira banksii	Acts as a chemoattractant	[71]
		Xiphophora chondrophylla
		Xiphophora gladiate
		Durvillaea potatorum
		Durvillaea antarctica
		Durvillaea willana
		Colpomenia peregrina		[72]
		Planosiphon complanatus (as Scytosiphon lomentaria)		[72]
		Analipus japonicus		[73]
	Ectocarpene	Ectocarpus flagelliformis (as Ectocarpus fasciculatus)	Acts as a chemoattractant	[71]
		Adenocystis longissima (as Adenocystis utricularis)
		Sphacelaria rigidula
		Analipus japonicus		[73]
	Dictyopterene C′	Dictyota dichotoma	Acts as a chemoattractant	[71]
	Dictyotene	Dictyota diemensis	Acts as an erotactin	[74]
	Desmarcstene	Desmarestia aculeata/Desmarestia menziesii (as Desmarestia aculeata)	Acts as a chemoattractant	[71]
		Desmarestia confervoides (as Desmarestia viridis)
		Cladostephus hirsutus/Cladostephus kuetzingii (as Cladostephus spongiosus)
	Lamoxirene	Laminariaceae	Acts as a chemoattractant	[71]
		Alariaceae
		Lessoniaceae
		Pleurophycus gardneri	Acts as an erotactin	[74]
		Agarum clathratum (as Agarum cribrosum)
		Sachharina gyrata (as Kjellmaniella gyrata)
		Hedophyllum sessile
		Cymathaere triplicate
		Undaria pinnatifida
		Pterygophora californica
		Eisenia bicyclis (as Eisenia arborea)
		Ecklonia biruncinata (as Ecklonia radiata)
		Macrocystis pyrifera
		Macrocystis pyrifera (as Macrocystis integrifolia)
		Nereocystis luetkeana
		Pelagophycus porra
		Dictyoneurum reticulatum (as Dictyoneuropsis reticulata)
		Lessoniopsis littoralis
	Multifidene cis-4-vinyl-5-cis- buten-l-yl-cyclopentene)	Cutleria multifida	Responsible for chemotaxis of the male microgametes	[75]
	Multifidene cis-4-vinyl-5-cis- buten-l-yl-cyclopentene)	Zonaria angustata	Acts as a chemoattractant	[73]
	Viridiene	Microzonia phinneyi (as Syringoderma phinneyi)		[71]
	Caudoxirene	Perithalia caudata		[71]
	Giffordene	Feldmannia mirchelliae/Hincksia mitchelliae (as Giffordia mitchelliae)		[72]
Rhodophyceae	Ochtodene (Monoterpene)	Ochtodes secundiramea	Acts as a deterrent to herbivores	[76]
Rhodophyceae	Octodiol	Ochtodes secundiramea	Acts as a deterrent to herbivores	[76]

Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

© 2023 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/).

Share and Cite

MDPI and ACS Style

Rathod, S.G.; Bhushan, S.; Mantri, V.A. Phytohormones and Pheromones in the Phycology Literature: Benchmarking of Data-Set and Developing Critical Tools of Biotechnological Implications for Commercial Aquaculture Industry. Phycology 2024, 4, 1-36. https://0-doi-org.brum.beds.ac.uk/10.3390/phycology4010001

AMA Style

Rathod SG, Bhushan S, Mantri VA. Phytohormones and Pheromones in the Phycology Literature: Benchmarking of Data-Set and Developing Critical Tools of Biotechnological Implications for Commercial Aquaculture Industry. Phycology. 2024; 4(1):1-36. https://0-doi-org.brum.beds.ac.uk/10.3390/phycology4010001

Chicago/Turabian Style

Rathod, Sachin G., Satej Bhushan, and Vaibhav A. Mantri. 2024. "Phytohormones and Pheromones in the Phycology Literature: Benchmarking of Data-Set and Developing Critical Tools of Biotechnological Implications for Commercial Aquaculture Industry" Phycology 4, no. 1: 1-36. https://0-doi-org.brum.beds.ac.uk/10.3390/phycology4010001

Article Menu

Phytohormones and Pheromones in the Phycology Literature: Benchmarking of Data-Set and Developing Critical Tools of Biotechnological Implications for Commercial Aquaculture Industry

Abstract

1. Introduction

2. Hormones and Pheromones Reported in the Phycology Literature

3. Biosynthetic Pathways

3.1. Hormones

3.1.1. Auxin

3.1.2. Cytokinins

3.1.3. Gibberellins

3.1.4. Abscisic Acid (ABA)

3.1.5. Ethylene

3.2. Pheromones

3.2.1. Giffordene

3.2.2. Dictyotene

3.2.3. Cystophorene

3.2.4. Ectocarpene

3.2.5. Dictyopterene

4. Role of Hormones in Algae

4.1. Auxin

4.2. Cytokinins

4.3. Gibberellins (GAs)

4.4. Abscisic Acid (ABA)

4.5. Ethylene

4.6. Brassinosteroids (BRs)

4.7. Jasmonic Acid (JA)

4.8. Polyamines (PAs)

4.9. Salicylic Acid (SA)

4.10. Strigolactone (SL)

4.11. Rhodomorphin

5. Role of Pheromones in Algae

5.1. Sporulation Inhibitors

5.2. Swarming Inhibitors

5.3. Ectocarpene

5.4. Dictyotene and C11 Sulfur Compounds

5.5. Ochtodene

5.6. Other Chemoattractants

6. Mode of Action of Hormones in Algae

7. Mode of Action of Pheromones in Algae

8. Methods for Extraction, Identification, and Quantification of Hormones from Algae

9. Methods for Extraction, Identification, and Quantification of Pheromones from Algae

10. Perspectives

11. Way Forward

Supplementary Materials

Author Contributions

Funding

Data Availability Statement

Acknowledgments

Conflicts of Interest

References

Share and Cite

Article Metrics

Article Access Statistics

Further Information

Guidelines

MDPI Initiatives

Follow MDPI