The enzymes choline acetyltransferases or choline acetylases (EC 2.3.1.6) participate in the synthesis of acetylcholine from choline and acetic acid (Nachmansohn and Machado 1943). A cell-free enzyme preparation with “choline acetylase” activity was present in Lactobacillus plantarum (Girvin and Stevenson 1954). This type of enzyme activity has also been found in many plant species (Roshchina 1991, 2001a).

Enzymes which degrade acetylcholine to choline and acetic acid are named cholinesterases and were first found in 1937 by Loewi in the hearts of amphibia. The function of acetylcholinesterase at cholinergic synapses of animals is to terminate cholinergic neurotransmission (Augustinsson 1949). However, the enzyme is expressed in tissues that are not directly innervated by cholinergic nerves. Moreover, transient expression in the brain during embryogenesis suggests that acetylcholinesterase may function in the regulation of neurite outgrowth. Overexpression of cholinesterases has also been correlated with tumorigenesis and abnormal mega-karyocytopoiesis (Small et al. 1996). Cholinesterase is also found in unicellular animals such as Paramecium (Corrado et al. 1999). An immunoblot analysis of the Paramecium enzyme revealed that the acetylcholinesterase had a molecular mass from 42 to 133 kDa, as reported for an analogous enzyme isolated from higher organisms. Structural homologies between cholinesterases and the adhesion proteins indicate that cholinesterases could also function as cell–cell or cell–substrate adhesion molecules. Abnormal expression of cholinesterases of both types has been detected around the amyloid plaques and neurofibrillary tangles in the brains of patients with Alzheimer’s disease (Small et al. 1996).

As for microorganisms, Goldstein and Goldstein (1953) first described the production of cholinesterase by a strain of bacterium Pseudomonas fluorescens after the culture was grown with acetylcholine as the sole source of carbon. The P. fluorescens enzyme was inducible, mainly, by choline (not as a carbon substrate, but, perhaps, as a source of nitrogen) or by two- to threefold less by some choline esters: acetylcholine > propionylcholine = benzoylcholine > butyrylcholine > acetyl-β-methylcholine. Addition of glucose completely prevented induction of the P. fluorescens enzyme. The pH optimum for growth of the culture and cholinesterase activity was 7.0, although the culture growth was higher in alkaline medium, where spontaneous hydrolysis of acetylcholine is also maximal. Choline oxidase synthesis in P. fluorescens has also been induced by choline. The cholinesterase of the bacterium may hydrolyze acetylcholine or propionylcholine, but to a lesser degree butyrylcholine, benzoylcholine, or acetyl-β-methylcholine. Like cholinesterase in animals, the enzyme activity in P. fluorescens was inhibited by neostigmine, with complete inactivation observed at high concentrations (10⁻³–10⁻² M) and only partly at the lower levels of 10⁻⁶ M. These levels of inhibition are similar to that observed in mammalian organ systems. The P. fluorescens protein was isolated and characterized (Goldstein 1959; Searle and Goldstein 1957, 1962; Fitch 1963a, b). Moreover, the strains of P. fluorescens tested exhibited preference for acetylcholine for growth promotion over choline, glycerol, glucose, succinate, betaine, and serine (Fitch 1963a). The isolated cholinesterase was inhibited by neostigmine in smaller (1000 times) concentration, than by physostigmine, but was not inhibited by diisopropylfluorophosphate (Fitch 1963b). A bell-shaped substrate saturation curve was observed, and specific activity of the 115-times purified cholinesterase was 10.5 mmol/mg protein/min. The enzyme had features both of true cholinesterase and acetylcholinesterase (Laing et al. 1967, 1969). Specific activity of the cholinesterase from P. fluorescens purified 40-fold by CM-50 Sephadex was up to 70 mmol/mg protein/min. The values of Km at pH 7.4 and 37 °C were 1.4 × 10⁻⁵ M for acetyl-choline and 2.0 × 10⁻⁵ M for propionylcholine, respectively, while butyrylcholine and benzoylcholine were not hydrolysable at all. The purified enzyme was inhibited by organophosphorus compounds and neostigmine, but not by physostigmine.

Imshenetskii et al. (1974) showed that a large variety of microorganisms may decompose acetylcholine including 31 strains of bacteria (genera Arthrobacter and Pseudomonas) and two strains of fungi (from 194 strains studied) that live in soil. Around 100–200 mg of wet biomass of active microbial strains were able to decompose 15–30 mmol of acetylcholine during a 2 h incubation, with the most active strains (50 mg of wet biomass) able to degrade up to 10 mmol/min. This active soil strain was identified as Arthrobacter simplex var. cholinesterasus var. nov. The amount of the decomposed acetylcholine by this microbe was 30 times higher than in other strains (Pseudomonas fluorescens—4 mM/h, P. aeruginosa—1 mM/h), while Arthrobacter simplex var. cholinesterasus var. nov had an activity of 300 mM/h. Actinomycetes (except two strains) and yeast had no significant cholinesterase activity. Our experiments showed a small rate of hydrolysis of acetylthiocholine and butyrylthiocholine in extracts of Saccharomyces cerevisiae where only 20–30 % inhibited by neostigmine or physostigmine, inhibitors of animal cholinesterase. Roshchina and Alexandrova (1991) isolated cholinesterase from the mycelium of the fungus Aspergillus niger. The enzyme hydrolyzed cholinic esters at rates similar with animal enzyme (K_M 7 × 10⁻⁴ M). Moreover, the enzyme has a molecular mass about 600 kDa determined by gel-filtration on Sephadex G-200 and after gel-electrophoresis with sodium dodecyl sulphate—two subunits 63 and 44 kDa. The enzyme activity was higher with acetylthiocholine, than butyrylthiocholine, and neostigmine inhibited the substrates’ hydrolysis, unlike physostigmine.

Cholinesterase activity has also been found in lower groups of the plant kingdom: in extracts of Characeae algae Nitella by Dettbarn, in 1962 and mycelium of fungi Physarum polycephalum by Nakajima and Hatano in 1962, and then a series of classical papers of Jaffe and Fluck with coworkers in 1970–1975 were devoted to the observation of the enzyme in many plant species: ~118 terrestrial species and ten marine algae were identified as having cholinesterase activity (for more details see the relevant references in monographs Roshchina 1991, 2001a). The values of the enzyme activity (the substrate hydrolysis rate) in higher plants were an average of 1–900 mmol/h/g fresh weight, depending on the plant species. It was shown that Bryophytes (mosses, liverworts and hornworts) demonstrate maximal cholinesterase activity of up to 0.360 mmol/h/g fresh weight (Gupta et al. 2001). Pollen also contain the enzyme (Bednarska and Tretyn 1989; Bednarska 1992; Roshchina et al. 1994; Rejon et al. 2012). Thus, detection of cholinesterase activity could serve as an additional indicator of the presence of acetylcholine.

Recently, the identification, purification, and cloning of maize acetylcholinesterase provided the first direct evidence of the enzyme formation in plants (Sagane et al. 2005). An especially important fact is that the acetylcholinesterase distribution in seedlings is sensitive to gravity, leading to asymmetry of the enzyme distribution (Momonoki 1997). Maize acetylcholinesterase activity localized in vascular bundles the coleoptile node and mesophyll seedlings is enhanced through a post-translational modification response to heat stress (Yamamoto and Momonoki 2012). The gene purportedly encoding for acetylcholinesterase was cloned from maize (Muralidharan et al. 2013). The Arabidopsis protein encoded by the gene exhibited lipase activity with preference for long chain substrates but did not hydrolyze choline esters. In several edible fruits and vegetables, members of the Solanaceae family (potato, eggplant and tomato), acetylcholinesterase inhibitors have been detected; these were mainly alkaloids which induced gastrointestinal and neurological symptoms in mammalian.

Plant cholinesterases are included in several processes (Roshchina and Semenova 1990; Roshchina 2001a, b), including but not limited to: pollen germination release just after moistening of the pollen grains (Roshchina et al. 1994; Roshchina 2007), as well as within the intercellular interactions in the pollen-pistil system (Kovaleva and Roshchina 1997). Cholinesterase may prevent the binding of dopamine with its receptor and decrease the toxic effect of d-tubocurarine (antagonist of nicotinic type of cholinoreceptor) on cells of horsetail vegetative microspores (Roshchina et al. 2012). Yamamoto and co-workers (2011) postulated that acetylcholinesterase is a positive regulator of heat tolerance in plants.

The biosynthetic pathway of biogenic amines includes the decarboxylation and hydroxylation of corresponding amino acids, in particular phenylalanine for the catecholamines, tryptophan for serotonin, and histidine for histamine (Lawrence 2004). Phenylalanine, precursor of dopamine, norepinephrine and epinephrine, is first hydroxylated, transforming to tyrosine and then to dihydroxyphenylalanine (DOPA). These processes are catalyzed by phenylalanine hydroxylase or phenylalanine monoxidase and tyrosine hydroxylase or tyrosine-3-monoxidase. Dopamine, an immediate precursor of norepinephrine and epinephrine, arises from DOPA through decarboxylation by means of the enzyme decarboxylase dioxyphenylalanine and the decarboxylase of aromatic amino acids (EC 4.1.1.26). Another route of tyrosine transformation is via decarboxylation, when it transforms to tyramine, and then by hydroxylation with the participation of tyramine hydroxylase into dopamine, which is then oxidized to norepinephrine by the copper-containing enzyme- β-hydroxylase 3,4-dioxyphenylethylamine. Then, under the influence of the transmethylase of phenylethanolamines, the formation of epinephrine takes place.

In the catabolism of catecholamines, aminooxidases participate as in the oxidative deamination of the catecholamines to metanephrine, normetanephrine, vanillic aldehyde, dehydroxymandelic and vanillic acids. For microorganisms, this metabolic process has not yet been studied. In plants, diamineoxidases play the main role in catecholamine metabolism, unlike animals that use monoaminooxidases for this purpose (Roshchina 2001a). As for catecholamine-O-methyltransferases, they are present in all animal tissues, and especially active in nervous cells. In plants, the catecholamine-O-methyltransferases pathway is also possible because the last three compounds are ordinary products of plant metabolism (Kuklin and Conger 1995; Roshchina 2001a; Kulma and Szopa 2007). Besides the abovementioned pathways of metabolism, catecholamines are oxidized by oxygen in air, forming oxidized products—red pigments aminochromes and black-brown pigments melanines which are polymers of indole (found both in plant and animals). The mechanism of oxidation is connected with the generation of superoxide radicals. Prevention of the oxidation of dopamine by superoxide dismutase confirms this mechanism. Enzymatic oxidation of catecholamines to melanines by polyphenol oxidase has been demonstrated (Roshchina 2001a). The abovementioned enzymes are found only in animals and plants. There is little data for catecholamine oxidation in microorganisms, although monoaminooxidase activity in mycobacteria (Pershin and Nesvadba 1963) and E. coli (Takenaka et al. 1997) has been found.

Serotonin is synthesized in plants and animals from tryptophan formed by the shikimate pathway, which has also been proposed for microorganisms (Oleskin et al. 1998a, b; Oleskin 2007). This process proceeds by two pathways: either via 5-hydroxytryptophan or tryptamine formation, or the first step of serotonin biosynthesis via decarboxylation of tryptophan, which then transforms in plants to tryptamine by action of the enzyme tryptophan decarboxylase (EC 4.1.1.27), or by the decarboxylation of aromatic amino acids (EC 4.1.1.26/27). Then, tryptamine is transformed to serotonin by hydroxylation with participation of the enzymes tryptamine-5-hydroxylase or 1-tryptophan-5-hydroxylase (EC 1.14.16.4). Hydroxylation of tryptophan leads to the formation of 5-oxytryptophan in the presence of tryptophan-5-hydroxylase (EC1.14.16.4). At the next stage, 5-oxytryptophan is decarboxylated by the decarboxylation of aromatic acids to yield serotonin. Tryptamine 5-hydroxylase, which converts tryptamine into serotonin and is common in animals, was also found as a soluble enzyme that had maximal activity in rice roots (Kang et al. 2007). The tissues of rice seedlings grown in the presence of tryptamine exhibited a dose-dependent increase in serotonin in parallel with enhanced enzyme activity. However, no significant increase in serotonin was observed in rice tissues grown in the presence of tryptophan, suggesting that tryptamine is a bottleneck intermediate substrate for serotonin synthesis. If we compare the enzymes from the different kingdoms, we can see more similarity. In particular, in the plant genus Arabidopsis, there is a homolog to part of a DNA binding complex corresponding to the animal tyrosine and tryptophan hydroxylases (Lu et al. 1992). Fujiwara et al. (2010) have found that unlike tryptophan hydroxylase of animals, an analogous enzyme in plants is a cytochrome P450 located in the membranes of the endoplasmic reticulum. The enzyme was not expressed in the bacteria E. coli, but may be expressed here via special genetic construction with recombinant genes (Park et al. 2011). Aminooxidases of biogenic amines may differ in microorganisms in relation to substrate specificity, in particular for the bacterium Methanosarcina barkeri and infusoria Tetrahymena pyriformis (Yagodina et al. 2000). Both studied enzymes can deaminate serotonin, but not histamine. The existence of one active centre for substrate binding is postulated as the aminooxidase of the bacterium, while several centres are thought to exist in the Infusorians.

For all living organisms, the biosynthesis pathway of histamine includes histidine decarboxylase which participates in the decarboxylation of histidine (Roshchina 2001a; Boron and Boulpaep 2005; Martín et al. 2005). The gene encoding histidine decarboxylase (hdcA) has been identified in different Gram-positive bacteria (Martín et al. 2005). Histidine decarboxylase used to be part of a cluster that included a gene of unknown function (hdcB) and a histidine–histamine antiporter gene (hdcC) in Pediococcus parvulus 276 and Lactobacillus hilgardii 321 has been identified (Landete et al. 2005). Catabolism of histamine occurs also via methylation or acetylation in the presence of histamine-N-methyltransferase, or histamine-N-acetyltransferase, and genes coding of the enzymes have been found in bacteria, plants, and animals (Iyer et al. 2004).

Appreciation of the presence and role of neurotransmitters in microbial cells was previously considered mainly in the context of receptors specific to the neurotransmitter compounds (Adler 1969), with particular understanding gained from concepts of neurotransmitter reception in animals. A major method by which organisms may be studied is the use of pharmacological assays wherein the neurotransmitter of a cellular reaction is analyzed via the use of agonists and antagonists to that neurotransmitter. All investigations concerning non-neuronal (non-synaptic) systems have a fundamental similarity to traditional studies undertaken for the nerve cell.

For acetylcholine, there are two types of acetylcholine receptors: nicotinic (receptors that respond to nicotine) and muscarinic (receptors that are sensitive to muscarine). Corrado et al. (2001) showed the presence of functionally related nicotinic and muscarinic receptors and the lytic enzyme acetylcholinesterase in the unicellular animal Paramecium primaurelia. In plants, the presence of similar receptors has also been shown (Roshchina 2001a). Recently, it was established that muscarinic and nicotinic acetylcholine receptors are involved in the regulation of stomata function—the opening and closing movement—in the plants Vicia faba and Pisum sativum (Wang et al. 1998, 1999a, 2000). Leng et al. (2000) showed the regulation role of acetylcholine and its antagonists in inward rectified K⁺ channels from Vicia faba guard cells. Location of the muscarinic receptor was shown in plasmatic membrane and chloroplast membranes (Meng et al. 2001), and cholinesterase activity was found in the cells (Wang et al. 1999b). The germination of plant microspores such as vegetative microspores of horsetail Equisetum arvense or pollen (generative microspores) of knight’s star Hippeastrum hybridum was blocked by the antagonists of acetylcholine, which are linked with nicotinic cholinoreceptors and Na⁺/K⁺ ion channels (Roshchina and Vikhlyantsev 2009). The nicotinic cholinoreceptors were cytochemically identified in the single-cell amoebae Dictyostelium discoideum, slugs, and spores, however, proteins immunologically-related to the muscarinic receptors were not present in the spores (Amaroli et al. 2003). Interestingly, nicotine and acetylcholine as the ligands of human nicotinic cholinoreceptors in culture of epithelial cells HEp-2 may stimulate the growth of Chlamydia pneumoniae (Yamaguchi et al. 2003).

The receptors for biogenic amines which are unique to animals are known as dopamine receptors, adrenoreceptors, and serotonin and histamine receptors. Similar receptors were observed in bacteria (Lyte and Ernst 1993; Lyte 2004; Freestone et al. 2007) and in plant cells (see monographs Roshchina 1991, 2001a). Alpha and beta adrenergic-like receptors may be involved in catecholamine-induced growth of Gram-negative bacteria (Lyte and Ernst 1993). In particular, Freestone et al. (2007) showed the blockade of catecholamine-induced growth of E. coli, Salmonella enterica, and Yersinia enterocolitica by adrenergic and dopaminergic receptor antagonists. In plants, adrenoreceptors participate in cytoplasm movement, ion permeability, and membrane potential, in flowering of Lemna paucicostata, photophosphorylation, as well as seeds, vegetative microspores’ and pollen germination (Roshchina 1991, 2001a; Baburina et al. 2000; Kulma and Szopa 2007). Serotonin- and histamine-sensitive receptors in plants regulate the seed, pollen, and vegetative microspores’ germination (Roshchina 1991, 2001a, 2004, 2005a). Shmukler et al. (2007) discussed earlier hypotheses of protosynapse for low-complexity organized animals and embryos of high-complexity organized animals, where the distribution of membrane serotonin receptors is restricted to the period of blastomere formation during cleavage and localized within the area of interblastomere contact. The hypothesis was based on their experiments where the membrane currents of the Paracentrotus lividus early embryos were registered after local application of serotonin drugs via special micropipette. Receptors of neurotransmitters may be linked with ion channels. Moreover, some domains of the ion channels appear connected to the cytoskeleton, in particular with actin (Cantiello 1997), and so the received chemosignal is likely to spread to the organelles via actomyosin filaments (Roshchina 2005a, 2006a, b).

Aspects of newer investigations of receptors are concerned with recombinant systems. For example, the recombinant mouse brain serotonin receptor (5HT1c) was used to study the response of plant cells and oocytes to a stress signal activated by the serotonin–serotonin receptor interaction and associated Ca²⁺ flux (Beljelarskaya and Sutton 2003). Based on plant expression vectors, recombinant constructs were obtained to direct production of 5HT1c fused with the green fluorescent protein in plant cells. The mRNAs for hybrid proteins were synthesized in an in vitro transcription system. The expression and function of the hybrid protein and the function of the associated ion channels were electrophysiologically studied in Xenopus laevis oocytes injected with the hybrid mRNA. The hybrid protein was functional and changed the operation of the Ca²⁺ channel in oocytes. To study the expression of the hybrid constructs in plant cells, the in vitro transcription product was inoculated in tobacco leaves, which thereupon fluoresced.

The function of compounds named neurotransmitters originates from simple chemotaxis and chemosignaling of microbial cells and leads to intercellular communication (Fig. 2.2). The so-called neurotransmitters may regulate (as hormones) growth and development of other unicellular organisms, and be attractants or repellents for them. In higher concentrations the same substances also play a role in defense (i.e. such as in the fight or flight response) or, in some cases, contribute to creating unique cultural foundations for food and cuisine. The following step in evolution includes the development of interactive relationships (parasitic, symbiotic, or otherwise) as well as the formation of multicellular organisms that may further specialize and attenuate the function of biomediators to serve unique requirements of communicative reaction-response within a larger, more complex multicellular system. This evolutionary pathway leads us to the concept of neurology as applicable to both animals and plants (Baluska et al. 2005; Brenner et al. 2006; Murch 2006; Ramakrishna et al. 2011).

In Table 2.3, the main functions of the neurotransmitters in all kingdoms are compared. The overall effect of a communication signal into a cell from an extracellular surface site or even intracellularly between cellular compartments, may occur via neurotransmitters. At the cellular level, the compounds may induce different reactions. Neurotransmitters are stored in secretory vesicles, and can be released inside or outside of the cell. Primary reaction to acetylcholine is often a change in membrane permeability for ions, while other reactions for both the neurotransmitter and biogenic amines are connected with the systems of secondary messengers—cyclic nucleotides, Ca²⁺, inositol-3-phosphate, and others.

A neurotransmitter functions as a chemosignal (i.e. the neurotransmitter is released from a cell and elicits a reaction in another cell) in the context of certain structures, including intercellular connections as well as intracellular environment such as between organelles within a cell. We can usually see the intercellular structural connection between plasma membranes of any cells that are in contact with one another, such as at the membranes of unicellular organisms or synaptic membranes of cells in organisms with a nervous system (Roshchina 1991, 2001a, b; Buznikov et al. 1996; Shmukler et al. 2007). As seen in Fig. 2.3, at any membrane point-of-contact either transitory or permanent, structural connections may be formed. There are junctions between endoplasmic reticulum and organelles within cells, or between different cells. Today, permanent or transitory structural-links between cells or within the cell are considered a necessary structural component in the transfer of the chemosignal.

As can be seen in Table 2.3, common cellular effects of neurotransmitters in living cells of all kingdoms include changes in membrane permeability (short-term effects) as well as the regulation of growth and development (long-term effects). The regulatory functions of neurotransmitters appear to have been conserved through evolution, effectively relating intracellular processes with unicellular populations within the environment. In this respect, the primary role of such compounds appear to be as a substrate in the acquisition of nitrogen by microorganisms (Imshenetskii et al. 1974), as well as serving as regulators in the microorganismal growth and development (Oleskin et al. 2010; Oleskin 2012).