Bioactive molecules from ciliates: Structure, activity, and applicative potential.

Ciliates are a rich source of molecules synthesized to socialize, compete ecologically, and interact with prey and predators. Their isolation from laboratory cultures is often straightforward, permitting the study of their mechanisms of action and their assessment for applied research. This review focuses on three classes of these bioactive molecules: (i) water-borne, cysteine-rich proteins that are used as signaling pheromones in self/nonself recognition phenomena; (ii) cell membrane-associated lipophilic terpenoids that are used in interspecies competitions for habitat colonization; (iii) cortical granule-associated molecules of various chemical nature that primarily serve offence/defense functions.

Bioactive molecules are generally recognize as those natural products—organic compounds that are synthesized by living systems—which common wisdom suggests to be structurally ‘optimized’ and selected for by evolution in order to confer their source organisms with a competitive, ecological or evolutionary advantage (Hanson, 2003). Bioactive molecules are commonly regarded as playing no direct role in the essential processes of the life. This role is distinctive of primary metabolites (Thirumurugan et al., 2018). Nevertheless, they govern and shape the reproductive success of their source organisms by driving defense/offence mechanisms, interactions with the environment and phenomena of self/nonself discrimination. This wide range of activities overall account well for the strong interest that bioactive molecules arouse both in basic research and bioprospecting for drug discovery and design in the fields of medical chemistry and pharmacognosy (Khazir et al., 2014; Newman & Cragg, 2014).

The vast array of effective sources of bioactive molecules is historically dominated by species from prokaryotes (Javed et al., 2011; Penesyan et al., 2010), plants (Chitra et al., 2019), (micro‐)fungi and filter‐feeding animals, in particular porifera, bryozoa and tunicates (Haefner, 2003; Kellner Filho et al., 2019; Molinski et al., 2009; Zhang et al., 2005). Species from the vast polyphyletic confederation of protists are largely relegated on the margins. However, research on easily cultivable diatoms, algae and ciliates provide evidence that also these microeukaryotes are a mine of natural products that deserve strong interest for biotechnological applications.

This article provides an overview of ciliate bioactive molecules that have been characterized to different extents for their chemical structures, analyzed for their activities and assessed for their potential applications. They are represented by (i) water‐borne signaling proteins collectively designated as ‘pheromones’, (ii) membrane lipid‐borne terpenoids collectively designated as ‘euplotins’ and (iii) cortical vesicle‐associated metabolites of more heterogeneous chemical nature (alkaloids, lactones and polyketides) variously named mostly with reference to their source ciliate.

WATER‐BORNE PHEROMONES

Ciliate pheromones are diffusible molecules that distinguish chemically conspecific cells, which are otherwise morphologically identical. In relation with their functional association with genetically determined systems of so‐called ‘mating types’—two or multiple within a species and questionably regarded as ‘sexes’—they regulate self/nonself recognition phenomena underlying the cell decision to shift from the vegetative (growth) life stage to the sexual stage manifested as cell–cell unions in mating pairs (Luporini & Miceli, 1986).

Pheromones have a long story, lately reviewed (Luporini, Alimenti, et al., 2016; Luporini, Pedrini, et al., 2016), in experimental ciliatology. Their original identification in the supernatant of Euplotes patella cultures via mating induction assays dates back nearly one century (Kimball, 1942). However, their isolation and chemical characterization were to be pioneered only decades later by Akio Miyake and collaborators in Blepharisma japonicum (Miyake, 1981). The two pheromones isolated from B. japonicum are unique in that their chemical nature differs strikingly. One, designated ‘gamone‐1’ (or ‘blepharmone’), is a highly unstable glycoprotein of 272 amino acids and six covalently linked sugars (Sugiura & Harumoto, 2001). The second, designated ‘gamone‐2 (or ‘blepharismone’), was instead determined to be a very stable tryptophan derivative, namely a calcium‐3‐(2‐formylamino‐5‐hydroxybenzoil) lactate (Miyake, 1981). In addition to differing in their chemical nature, gamone‐1 and gamone‐2 also differ in their spectrum of specificity, only the former being species‐specific and the latter, instead, common to multiple species of Blepharisma (Kobayashi et al., 2015). This lack of species‐specificity, together with the eccentricity of the chemical nature have thus suggested that gamone‐2 may act not as a genuine pheromone, but rather as a hapten‐like molecule that binds to, and activates the glycoprotein gamone‐1 (Luporini & Miceli, 1986; Nanney, 1980). Support for this view derives from observing that B. japonicum, alike heterotriches in general, regularly practices self‐mating, which clearly implies that both gamones 1 and 2 (yet isolated from transiently not self‐mating cultures of two supposed distinct cell types, I and II) must be co‐released by the same cells.

Differently from Blepharisma pheromones, those isolated from Dileptus anser and various species of Euplotes, namely E. raikovi, E. octocarinatus, E. nobilii, E. petzi and E. crassus, all are high‐stable proteins that maintain long‐lasting integrity and biological activity in their natural environment. For the D. anser pheromones (three isolated from interbreeding cultures), the characterization is narrowed to a molecular mass in the range of 3–4.5 kDa and a strong propensity to associate into unstable oligomers (Parfenova et al., 1989; Uspenskaya & Yudin, 2016). Far more characterized are Euplotes pheromones, known to be encoded by multiple series of codominant alleles at the mat‐locus of the micronuclear genome (Vallesi et al., 2014). Their largely variable amino acid sequences, determined in significant numbers from each species, range in length from 32 amino acid residues in E. petzi (Pedrini et al., 2017) to 109 in E. octocarinatus (Brünen‐Nieweler et al., 1998), and limit an extensive intraspecific and interspecific conservation only to the cysteine residues that are regularly paired into intrachain disulfide bonds (Figure 1). Their variability contrasts with the remarkable extent of intraspecific and interspecific identity of the signal/prepeptide and pro‐segment sequences that are proteolytically cleaved from the immature cytoplasmic pheromone precursors (pre‐pro‐pheromones) before the secretion of the active pheromone (Ricci et al., 2019). Variations in the pheromone amino acid sequences may also closely reflect deep interspecies differences in ecological adaptation. Instructive is the case of the cold‐adapted pheromones from the bipolar species, E. nobilii, which unfold in the range 55__70°C, as compared to their homologs from the closely related temperate‐water species, E. raikovi, that unfold only over 95°C (Cazzolli et al., 2013; Geralt et al., 2013). Like psychrophilic proteins, E. nobilii pheromones are significantly richer than E. raikovi pheromones in polar amino acids (44% vs. 30%), in particular threonine (12% vs. 6%) and asparagine (8% vs. 4%), and poorer (44% vs. 57%) in hydrophobic ones, in particular leucine (1% vs. 7%) and isoleucine (2% vs. 6%). These chemical features are overall consistent with stronger molecular interactions with the solvent and weaker stabilizing forces of the protein core (Alimenti et al., 2009).

FIGURE 1

Amino acid sequences and NMR solution structures of Euplotes pheromones. (A) For each species‐specific pheromone family, represented by the members listed between brackets, only the consensus sequence is reported. The conserved cysteines are highlighted in red, and X stands for any amino acid. The sequence of the E. crassus pheromone Ec‐α, the only one known from a sub‐family distinct from that which includes the other pheromone sequences, is reported individually and written in the single letter code. Analogously, the sequences of the E. octocarinatus pheromone Phr4 and E. raikovi pheromone Er‐23 are reported individually, as they represent structurally deviant family members. The molecular structures of E. raikovi pheromones Er‐1 (PDB code, 1ERC) and Er‐23 (PDB code, 1HA8), E. nobilii pheromone En‐6 (PDB code, 2JMS), and E. petzi pheromone Ep‐1 (PDB code, 2N2S) are shown (in ribbon diagrams) as representative of each pheromone family. The disulfide bonds stabilizing the molecular structures are shown as yellow ball‐and‐stick diagrams. The helix labeled h3 in the Er‐1 pheromone structure is highlighted in red to indicate the tight intraspecific and interspecific conservation of the structural backbone. N and C identify the molecule amino and carboxyl termini, respectively. (B) Very simplified version of the Euplotes phylogenetic tree, articulated into six clades (boxes), showing the positions of the five species analyzed for the pheromone structures

By applying NMR spectroscopy and/or X‐ray crystallography to native protein preparations, a variety of Euplotes pheromone molecular structures have been resolved: Er‐1, Er‐2, Er‐10, Er‐11, Er‐13, Er‐22 and Er‐23 from E. raikovi; En‐1, En‐2 and En‐6 from E. nobilii; and Ep‐1 from for E. petzi. Comparative analyses of the specificities of these pheromone structures (the only known for water‐borne proteins from protists) were object of previous reviews (Luporini, Alimenti, et al., 2016; Luporini, Pedrini, et al., 2016). In this context, one most relevant concept is worth being reiterated in relation to the pheromone mechanism of action. Despite considerable variations in the amino acid sequences, Euplotes pheromones form species‐specific families of structurally homologous proteins that closely mimic one another in their molecular structures dictated by a common up‐down‐up three‐helix fold tightly stabilized by densely spaced disulfide bonds. In close functional analogy with animal cytokines and protein growth factors, they can thus elicit different and context‐dependent cell growth, or mating responses by competitively binding in autocrine (self), or heterologous (nonself) fashion to their receptors on the surface of target cells.

Insights into the molecular mechanisms underlying these competitive pheromone/receptor binding interactions derive from the study of the protein–protein interactions that arrange E. raikovi pheromone molecules into crystals (Pedrini et al., 2021; Weiss et al., 1995). The rationale for this study resides in the identification of the pheromone receptors with membrane‐bound pheromone isoforms (represented by type‐II transmembrane proteins oriented with the N‐ and C‐terminus on the cell inside and outside, respectively), that are specified through an intron‐splicing mechanism by the same macronuclear genes which encode the soluble pheromones (Ortenzi et al., 2000; Ricci et al., 2019, 2021). Due to this common genetic determination, in each cell type the extracellular domain of the pheromone isoforms comes to be structurally identical with the soluble pheromone, and the pheromone/ligand‐binding capability of this domain is supported by mass‐spectrometry evidence that native pheromone molecules can dimerize (or possibly form higher aggregates) with the subunits unstably associated by noncovalent forces (Bradshaw et al., 1990).

Two lines of evidence emerge by comparing the crystal structures of the E. raikovi pheromones Er‐1 and Er‐13 (Pedrini et al., 2021; Weiss et al., 1995), that are specific of strongly mating compatible cell types (I and XIII, respectively) and show largely overlapping backbone molecular structures (Figure 2). First, Er‐1 and Er‐13 molecules interact similarly in a cooperative fashion to tightly associate into linear chains that propagate along a two‐fold screw‐symmetry rotation axis and include subunits that are rigorously ordered with mutually opposite orientations as expected to be the case between pheromone and receptor moieties interacting on a cell surface. Second, although Er‐1 and Er‐13 molecules take different intrachain arrangements, which reflect a marked difference in the symmetry space group of the Er‐1 and Er‐13 crystals (C 2 and P43 groups, respectively), both mostly use residue side chains lying on the third one of the three helices of the basic pheromone molecular scaffold to establish the direct contacts which are responsible for the widest intrachain contact interface.

FIGURE 2

Crystal structures of E. raikovi pheromones Er‐1 and Er‐13. (A) Amino acid sequences and molecular structures of the two pheromones. In the amino acid sequences, the three helical segments (h1, h2 and h3) are boxed, and the six cysteine residues are indicated by progressive Roman numerals and connected according to their disulfide pairings represented by yellow ball‐and‐stick diagrams. Helix 3 is highlighted in red to indicate the central role played in establishing the contacts that pack molecules into crystals. (B) Perspective view (on the left) and top view (on the right) of the one‐dimensional linear chains, propagating along a two‐fold screw‐symmetry rotation axis (red symbol), that Er‐1 and Er‐13 molecules equally form in the crystal taking rigorously alternating up and down orientations, as indicated by the positions of the amino (N) and carboxyl (C) termini. The Er‐1 and Er‐13 chains include a common protein–protein contact interface (green bars), that is equally derived from burying amino acid side‐chains mostly residing on h3. However, they differ in the geometrical arrangement of the molecules, as denoted by observing that Er‐1 molecules expose both h1 and h2 parallel to the chain sides, while Er‐13 molecules expose only h2. In the Er‐1 crystal (PDBs, 1ERL, 2ERL and 6E6O), this difference reflects in a further side‐by‐side chain association into bidimensional layers (not shown) and in two‐fold symmetries specific of the space group C2. In the Er‐13 crystal (PDB, 6E6N), it reflects in a mutual arrangement of the chains at 90° between one another (not shown), which prevents the layer formation and is consistent with four‐fold screw symmetries distinctive of the space group P43

Helix‐3 is thus identified as the key functional player in the pheromone/receptor interactions, and this view is strongly supported by biological and phylogenetic observations. On biological grounds, the most significant observation is relevant to a change in the pheromone activity that is observed in aging E. raikovi cultures and caused by the oxidation of a Met‐residue lying exposed on the helix‐3 of the pheromone Er‐1 (Alimenti et al., 2012). Oxidized Er‐1 molecules bind to their source type‐I cells no longer as an autocrine growth‐promoting signal, but as a heterologous (nonself) signal that induces homotypic (self) mating pair formation between their source cells. On phylogenetic grounds, the overall comparison of the E. raikovi, E. nobilii and E. petzi pheromone molecular structures clearly indicates that helix‐3 is much more tightly conserved than the two other helices in relation to the backbone trace, extension and regular alpha‐conformation. At intraspecific level, its conservation also extends to the structurally eccentric E. raikovi pheromone Er‐23, in which the three‐helix bundle is substantially changed into a five‐helix bundle (see Figure 1) by the accommodation of two supplementary helical turns in consequence of the incorporation of a new segment of 11 residues (for a total of 51) and four new cysteine residues (for a total of 10; Zahn et al., 2001). At interspecific level, it finds a close counterpart with the largely predominant helix of E. petzi pheromones (see Figure 1), that may be considered as the evolutionary forerunners of all the other Euplotes pheromones in the light of the basal branch that the E. petzi clade forms in the Euplotes phylogenetic tree (Di Giuseppe et al., 2014). Lastly, the likely general helix‐3 conservation among Euplotes pheromones may well explain the numerous cases of pheromone‐mediated cross‐mating reactions that are observed by mixing heterospecific cell cultures (Kuhlmann & Sato, 1993; Nobili et al., 1978).

In studying functional similarities of Euplotes pheromones with mammalian protein growth factors, the immune‐stimulatory cytokine interleukin‐2 (IL‐2) was serendipitously found (along with epidermal growth factor and other signaling proteins) to act as a very effective competitor of pheromone/receptor binding reactions (Ortenzi et al., 1990). The significance of this finding is reinforced by observing that the E. raikovi pheromone Er‐1 binds with different affinities to the α and β chains of the trimeric IL‐2 receptor on the IL‐2‐dependent mouse T lymphocyte CTLL‐2 cells, and increases (at nanomolar concentrations) DNA synthesis, proliferation and viability of human lymphoid Jurkat T‐cells (Vallesi et al., 1998). In these cells, it also triggers the mitogen‐activated protein kinase ERK1/2 pathway, increases the expression of an array of cytokines (including interferon‐γ, tumor necrosis factor‐α, IL‐1β, IL‐2 and IL‐13), and induces the synthesis and release of factors inhibiting the cell cycle progression of human glioma U‐373 cells (Cervia et al., 2013).

MEMBRANE‐ASSOCIATED EUPLOTINS

The term ‘euplotin’ should strictly (and correctly) be applied to the earliest set of sesquiterpenoids that have been characterized from Euplotes crassus strains (Guella et al., 1994). It is here used in the plural form to collectively designate all the terpenoids that mainly localize into the membrane lipid bilayer of marine species of Euplotes, and have then been purified from large‐scale cultures by liquid chromatography and structurally resolved by advanced analytical techniques such as mass spectrometry and nuclear magnetic resonance spectroscopy.

The euplotin terpenoid structures vary remarkably between species, and may to some extent be credited as species‐specific chemo‐taxonomic markers. Despite their wide chemo‐diversity, all originate from two distinct enantiomeric biogenetic precursors, farnesyl or geranyl‐geraniol pyrophosphates, through biosynthetic pathways leading to sesquiterpenoid and diterpenoid structures, respectively, which have mostly been named in relation to their source species and grouped into four distinct classes in relation to their structural backbone (Figure 3).

FIGURE 3

The four distinct classes of chemical structures of lipophilic terpenoids, designated euplotins, characterized from different marine species of Euplotes. Arrows indicate biosynthetic derivatives from precursor forms

The first class, built on an unsaturated dioxa‐tricyclic backbone, includes three E. crassus‐specific sesquiterpenoid euplotins named ‘euplotin A’, ‘euplotin B’ and ‘euplotin C’ (which is systematically detected from all the strains as the most abundant compound) plus the common biogenetic acyclic precursor, ‘preuplotin’. The proposed biosynthetic route relies on the oxidation of the methyl groups of the farnesyl pyrophosphate that leads to the acyclic precursor preuplotin followed by an unexpected internal cyclization leading to the euplotin skeleton (Guella et al., 1994).

The second class includes two terpenoid euplotins containing a polycyclic structure built on a novel C‐30 backbone, isolated from some (tropical) E. vannus strains and named ‘vannusal A’ and ‘vannusal B’, plus three sesquiterpenoid euplotins isolated from other (temperate‐water) conspecific strains and named ‘hemivannusal’, ‘prevannusadial A’ and ‘prevannusadial B’ in relation to their likely role as vannusal A and B precursors (Guella et al., 2007).

The third class includes sesquiterpenoid euplotins isolated from E. rariseta strains (Guella et al., 1996a) and diterpenoid euplotins isolated from E. focardii (Guella et al., 1996b) that are similarly built on an octahydroazulene ring system. In the E. rariseta case, strains representing two morphologically and genetically distinct populations synthesize two epimeric forms of which one, named ‘rarisetenolide’, is isolated together with its epoxide derivative, ‘epoxyrarisetenolide’, and the second, named ‘epirarisetenolide’, is the corresponding C(10) epimer. The two epimeric forms may equally be derived from alternative enzymatic biosynthetic pathways, or two distinct enantiomeric biogenetic precursors. In any case, the enzymatic steps resulting in the construction of the octahydroazulene ring system of rarisetenolide and epirarisetenolide appears to be a specific feature of the ciliate secondary metabolism. In the case of E. focardii (represented by a single strain endemic to Antarctic waters), two diterpenoid euplotins, named ‘1‐S‐focardin’ and ‘1‐R‐focardin’, are isolated as an equilibrating mixture of two diasteroisomeric hemiacetals. They are in equilibrium with the unstable ring‐opened aldehyde intermediate ‘prefocardin’, and accompany with ‘epoxyfocardin’ as a minor metabolite.

The last class is represented by a sesquiterpenoid euplotin built on a bicyclo[3.2.0]heptane ring system, that is specific to E. raikovi and named ‘raikovenal’ (Guella et al., 1995). As in E. rariseta, different E. raikovi populations differ in the production of C(10)‐epimeric forms. Some strains synthesize raikovenal together with its putative biogenetic precursor, ‘preraikovenal’, while others synthesize the C(10) epimer, ‘epiraikovenal’, together with its seco‐analog, ‘secoepiraikovenal’. Considering that a metabolite and its epimer require different biogenetic pathways, the production of functional alternatives embedded in the same bicyclo[3.2.0]heptane skeleton may be viewed as an ecological adaptation strategy.

In the natural environment, a major euplotin function is most likely played in habitat colonization by inhibiting via cell‐to‐cell contacts the growth of competitors for space and energy resources (Dini et al., 1993). Experimental support to this view was principally derived from using native preparations of euplotin C on cultures of one E. vannus strain, chosen as an ecological competitor to E. crassus populations and selected as one of the poorest synthesizers of vannusal A and B. At a concentration of 10 μg/ml, euplotin C is lethal in short time, while at minor sublethal concentrations it alters the competitor's cell cycle, ciliary motility and the body shape by compromising the electrical properties of the membranes. A few minutes of exposure are sufficient to impair the competitor ion channel pumps and cause a rapid increase in intracellular concentrations of both Ca2+ and Na+, which in turn threaten the cell motility (Trielli et al., 2008). Additionally, euplotin C decreases mitochondrial functions activating a caspase‐dependent type of apoptosis, enhances lysosomal pH, and reduces lysosomal membrane stability. Overall, these events result in the dismantling ciliary structures and microtubules, the development of aberrant cytoplasmic vacuoles, and a drop in phagocytic activity.

On other cell systems, the noxious effects of euplotin C extend from pathogenic microorganisms (such as Leishmania major, Candida albicans and several strains of Streptococcus and Burkholderia spp.) (Savoia et al., 2004), to various mammalian tumor‐derived cell lines. In corticotropic mouse pituitary tumor cells and rat pheochromocytoma cells, euplotin C induces a rapid activation of ryanodine receptors (a class of intracellular Ca2+ channels), a depletion of Ca2+ stores in the endoplasmic reticulum, the release of cytochrome c from mitochondria, and the caspase‐12/caspase‐3 activation which, in turn, triggers the apoptotic cell death pathway (Cervia et al., 2006, 2007). Assayed in comparison with cisplatin (a common chemotherapeutic agent used on many cancer types) on human melanoma cell lines, euplotin C similarly shows a good tolerability profile in significantly decreasing the melanoma cell viability and inhibiting the mitogen‐activated protein kinase ERK1/2 and protein kinase B/Akt signaling pathways, which control many aspects of the melanoma aggressiveness and cell migration (Carpi et al., 2015, 2018).

CORTICAL GRANULE‐ASSOCIATED BIOACTIVE MOLECULES

To interact with prey and predators, ciliates rely on very effective offense/defense mechanisms operating by means of an ample array of structurally diverse molecules that localize into extrusomes, or other nonextrusive (either pigmented or colorless) membrane‐bounded cortical organelles. These molecules, noxious to prey and/or predators, are mostly derived from various biogenetic precursors (acetate, shikimic acid, mevalonic acid and amino acids) of the primary metabolism, and have motivated particular interest from an applied perspective for chemical structures readier to be synthesized.

Previously studied for a function primarily played only in photoreception (Checcucci et al., 1997; Maeda et al., 1997; Tao et al., 1993), ‘blepharismins’ and ‘stentorin’__the pigment molecules that reside in the cortical granules of species of Blepharisma and Stentor and are structurally closely related to the naphthodianthrone hypericin native to the flowering plant, Hypericum perforatum __were later discovered to be in fact multifunctional (Lobban et al., 2007) and active in prey/predator interactions (Harumoto et al., 1998; Miyake et al., 2001). They were also utilized as antibiotics against the methicillin‐resistant Gram‐positive Staphylococcus aureus and the Gram‐negative Bacillus subtilis and B. cereus, and as cytostatic compounds on HeLa cells (Cavaleiro et al., 2020; Pant et al., 1997).

In the wake of these findings, new metabolites associated with pigmented or colorless cortical granules have been characterized for their defense/offense activities and chemical structures. Among them: ‘keronopsins’, ‘kerenopsamides’ and ‘erythrolactones’ (represented by three active nonsulfonated forms, A2, B2 and C2, laying in the cortical granules, and the respective inactive sulfonate esters in the cell cytoplasm) from Pseudokeronopsis rubra, P. riccii and P. erythrina, respectively (Anesi et al., 2016; Buonanno et al., 2017; Guella, Frassanito, et al., 2010; Höfle et al., 1994); ‘spirostomin’ and ‘mono‐prenyl hydroquinone’ from Spirostomum teres and S. amibiguum, respectively (Buonanno et al., 2012); and ‘climacostol’ from Climacostomum virens (Masaki et al., 1999; Miyake et al., 2003). The chemical structures of these molecules serving primary defense/offense functions are reported in Figure 4. Keronopsamides are alkaloids with a fully planar structure where the molecular skeleton is formally derived from a peptide‐like condensation of a 3‐pyrrolepropenoic acid with an unsaturated bromotyramine, while it is suggested that keronopsins and erythrolactones derive from the metabolism of fatty acid (Guella, Skropeta, et al., 2010). In the former, a β‐bromide‐substituted pyrrole is linked to a sulfate pyrrole through a conjugated acyl chain; in the latter, the 4‐hydroxy‐unsaturated d‐lactone ring is 2,6‐dialkyl substituted with saturated alkyl chain at carbon‐2 and a butenyl‐hydroquinone group at carbon‐5. More marked is the interspecific structural divergence between spirostomin and mono‐prenyl hydroquinone isolated from the two Spirostomum species (Buonanno et al., 2012), the former being derived from intramolecular cyclization of a C14 linear polyketide backbone and the latter being a prenylated‐hydroquinone derivative, a rather common metabolite among both unicellular and multicellular organisms. Lastly, climacostol is characterized by a resorcinol core to which an unsaturated C9‐alkenyl chain is attached (Masaki et al., 1999).

FIGURE 4

Bioactive molecules of different chemical nature from cortical granules of species of Pseudokeronopsis, Spirostomum and Climacostomum. In the three lactone molecules, different side chains attached to a same carbon skeleton are indicated by R1 and R2. In the three synthetic derivatives (indicated by arrows and named AN1, AN2 and MOMO) of native climacostol, the side chain modifications are in bold

In an applied perspective, climacostol has attracted much more interest than the other molecules in relation to two major experimental reasons: (i) the availability of substantial amounts of native preparations that have initially greatly facilitated antimicrobial and cytotoxic bioassays on a large spectrum of cell systems; and (ii) the concrete possibility to synthesize climacostol in both the native and modified more bioactive structures. Among these, one, characterized by a Z‐configuration of the C=C double bond at the 2′‐position of the carbon chain (Fiorini et al., 2010; Masaki et al., 1999), is also sold as apoptotic agent, available either as ‘cold’ or ‘deuterium‐labeled’ molecule (Santa Cruz Biotechnology, Dallas, USA; Advanced Technology & Industrial Co., Hong Kong, China).

The activity of climacostol as pharmacological drug has mostly been assayed on human lines of melanoma, promyelocytic leukemia and tumor squamous carcinoma cells. More reliable effects are recorded on melanoma cells in which climacostol rapidly causes (at micromolar concentrations) dysfunctional autophagy, nuclear and mitochondrial DNA damage with consequent activation of the caspase apoptotic pathway. The significance of these observations is further supported by in vivo experiments, in which the development of mouse melanoma allografts is halted at various extents of effectiveness by intratumor climacostol injections (Buonanno et al., 2020; Catalani et al., 2016).

The addition of a methyl or hydroxyl group to the aromatic ring of climacostol results into synthetic analogues capable to effectively modulate both potency and mechanism of action of the native molecule (Buonanno et al., 2019), while adding a methoxymethyl ether protective group to the aromatic ring results in the synthesis of an inactive prodrug, which changes to active climacostol only at pH < 7 (Catalani et al., 2019). Delivered to pathological cell systems, this prodrug has shown to be particularly effective on melanoma cells grown in mild acidic culture medium (pH 6.3), and appears to be promising against Leishmania and Plasmodium. In the former case, by acting in the acidic environment of the parasitophorous vacuole in the host's macrophages, in the latter, by acting in the acidic extracellular medium surrounding the infected red blood cells.

CONCLUDING REMARKS

Ciliates count a few equals among eukaryotic microorganisms in outcompeting for habitat colonization and energetic resources, and the utilization of a huge reserve of structurally and functionally diverse bioactive molecules accounts well for their success in facing natural selection. Although these molecules have been isolated and studied for their native and applied activities only from a few, more easily collected species, the observations surveyed here provide good evidence for deserving strong research interest. In basic science, to trace the evolutionary forerunners of the structures and mechanisms of action of the molecules that regulate the multicellular life; in applied science, to discover new lead, bio‐friendly compounds that, either in the native or a derived structure, may serve the fields of medicine and pharmacology.

In addition to plants (Lautié et al., 2020), marine fauna has also significantly contributed at the discovery of very effective chemo‐therapeutic agents (Khazir et al., 2014; Newman & Cragg, 2014). Common examples are cytarabine and halichondrin (the former used in various forms of leukemia and the latter in breast cancer) from species of the sponges Cryptotheca and Halichondria, respectively, or trabectedin (used in the soft‐tissue sarcoma and ovarian cancer treatment) from species of the tunicate Ecteinacidia (D’Incalci et al., 2014; Schwartsmann et al., 2001; Sipkema et al., 2005). However, the exploitation of the marine animal biodiversity as a direct source of pharmacologically utilizable molecules is often challenged by low yields and complex chemical structures, and may turn out to be detrimental to the ecological niche of a species and, in the longer term, result in ecosystem unbalance. By exploiting ciliates and other micro‐eukaryotes as well, the environmental impact is neutralized by the real possibility to grow large‐scale cultures of many species in captivity, with greatly reduced costs, no legal restrictions and no waste.

Euplotes species certainly figure among those, which are more easily collected and cultivated. In addition, they are unique for unmatched levels of interspecific and intraspecific genetic polymorphism arising from the evolution of high‐multiple mating‐type systems and a consequent acquisition of strongly outbreeding strategies (Nobili et al., 1978). In the case of euplotins, this polymorphism reflects as a wide diversification of skeletal structures requiring enzymatic pathways unorthodox with respect to the pathway common to the linear terpenoid precursors; in the case of pheromones, it results in the generation of an unrestricted number of physical–chemical specificities that superimpose on, and make unique a substantially common three‐helix protein fold. It also may provide practical advantages in identifying species and strains that are better able to experimentally satisfy quantitative needs in assaying the euplotin and pheromone activities. Among euplotins, up to 1 mg of euplotin C can be extracted from 1 ml of a cell pellet of certain E. crassus strains (Savoia et al., 2004); among pheromones, up to 300 µg of pure Er‐1 protein can be purified from 1 L of supernatant of the E. raikovi type‐I cell cultures (Raffioni et al., 1992). At the opposite end of the Euplotes species, Climacostomum virens, source of climacostol, is rarely collected and difficult to cultivate. In a sense, it compensates by producing a bioactive molecule that, in addition to looking as most promising in an applied perspective, has shown to be the best‐predisposed to structurally engineering and synthesis.