Flatworm nervous systems comprise central and peripheral components that facilitate coordinated and complex behaviors that are modulated by physiological status and sensory input. The absence of a body cavity in flatworms enhances their dependence on neuronal signaling for intercellular communication. Significant advances have been made in our understanding of the neurobiology of flatworms, largely through the growth in genomic/transcriptomic resources and some progress in the development of functional genomics tools. This chapter describes the "state of the art" of flatworm neurobiology with a primary focus on the recent advances made in parasitic flatworms where progress has been driven by the search for new targets for chemotherapeutics.
For several decades, helminth neurotransmission has been recognized as an appealing source of targets for new anthelmintic drugs (Geary et al., 1999; Maule et al., 2002; Mousley et al., 2004; Ribeiro et al., 2005). This appeal stems from two factors: first, the demonstrable importance of the neuromuscular system (NMS) in essential functions such as motility, chemosensation, reproduction, attachment, and feeding; second, many of the core receptors, enzymes, and transporters involved in neurotransmitter biosynthesis and signal transduction are demonstrably "druggable." That is, they represent protein families against which active compounds already exist in human and/or veterinary medicine. G protein–coupled receptors (GPCRs), ligand-gated ion channels (LGICs), and neurotransmitter transporters are particularly good examples of such targets.
This chapter aims to describe the current state of knowledge regarding the neurotransmitters, receptors, and signaling proteins employed by the flatworm nervous system, in the context of how their structural and functional data can be translated to novel control options for parasites.
In covering the entirety of flatworm neurotransmission, this is necessarily a broad review. Readers interested in more detail on specific areas are directed to reviews on classical transmitters (Ribeiro et al., 2005; Ribeiro, 2015), neurotransmitter transporters (Ribeiro & Patocka, 2013a), neuropeptides (McVeigh et al., 2005b; Marks & Maule, 2010), and anthelmintic drug discovery (Geary, 2010; Geary et al., 2015).
All four classes of phylum Platyhelminthes (Cestoda, Monogenea, Planaria, and Trematoda) display a common neuronal architecture, comprising central and peripheral nervous systems (the definitive description of flatworm neuroanatomy can be found in Halton & Gustafsson, 1996). Typically, the central nervous system (CNS) consists of a relatively primitive anterior brain, composed of paired anterio-lateral ganglia, from which neurons project posteriorly along the paired main nerve cords, laterally across the cerebral commissure, and anteriorly toward the mouth/oral sucker/anterior sense organs. Posterior to the brain, the longitudinal nerve cords are intermittently connected by transverse ring commissures to generate a distinctive orthogonal (ladder-like) appearance (Halton & Gustafsson, 1996). Peripheral motor and sensory neurons (peripheral nervous system, PNS) branch from the CNS, forming fine nerve plexuses that innervate the somatic muscle and surface sensory organs. The nerve cells themselves encompass unipolar, bipolar, and multipolar cells; are typically unmyelinated; and are often found in close association with what are interpreted to be glial cells (Halton & Gustafsson, 1996). Flatworm neurons interact with other cells at synapses that exhibit broad structural diversity. Typical synaptocrine chemical synapses include two-way (nonpolarized) junctions, as well as polarized synapses via which neurons may interact with single or multiple postsynaptic cells. In the case of neuromuscular junctions, nerves may make a traditional synapse onto muscle, or the muscle can reach out to the axon via a cytoplasmic extension. Nonsynaptic release sites exist, where neurosecretory vesicles may be exocytosed from the neuronal membrane directly onto an abutting nerve fiber, fulfiling a neuromodulatory role rather than that of a direct synaptic transmitter.
Flatworms employ classical/small molecule and peptidergic neurotransmitters, both of which are widely distributed throughout the CNS and PNS (Ribeiro et al., 2005; McVeigh et al., 2005b). These two neurotransmitter classes are generated by distinct biosynthetic means, but their synaptic release, receptor interactions, and signaling pathways employ relatively similar mechanisms. Classical transmitters are produced by biosynthetic enzymes present in the cytosol of a presynaptic nerve terminal, with each transmitter synthesized by a discrete set of enzymes (Fig. 1), before being loaded into secretory vesicles by transmitter-specific transporter pumps. Neuropeptides are produced via the protein synthesis pathway, as single or multiple copies of related peptides within larger precursor proteins (prepropeptides). Mature peptides are subsequently excised from the prepropeptide, ahead of their posttranslational processing. This processing occurs entirely within the secretory pathway of the endoplasmic reticulum (ER) and Golgi, resulting in vesicles filled with mature, processed peptides. Secretory vesicles of either classical or peptidergic origin accumulate at the presynaptic membrane, ready for release upon receipt of an appropriate signal. Generally, the arrival of an electrical impulse triggers the influx of calcium (Ca2+) into the cytosol through voltage-operated Ca2+ channels (VOCCs). Subsequently increased cytosolic [Ca2+] causes conformational changes in SNARE-like docking proteins (Hong & Lev, 2014), resulting in membrane fusion and vesicular exocytosis. Transmitters released into the synapse interact with receptors present on the postsynaptic membrane. Flatworm neurotransmitters most typically employ one of two types of receptors—ligand-gated ion channels (LGICs) and G protein–coupled receptors (GPCRs).
Figure 1 Biosynthetic pathways of classical neurotransmitters. Precursors and enzymes responsible for biosynthesis of the classical transmitters present in flatworms are shown. Accession numbers are presented for enzymes that have been cloned or reported in genome publications. RNA interference (RNAi) phenotypes are also summarized where available. References: DjChAT (Nishimura et al., 2010); Tyrosine hydroxylase (Hamdan & Ribeiro, 1998; Hu et al., 2011); DjGAD (Nishimura et al., 2008a); DjTBH (Nishimura et al., 2008b); and SmTPH (Hamdan & Ribeiro, 1999).
Among classical neurotransmitters there are examples of both LGIC- and GPCR- mediated signal transduction, while all the flatworm neuropeptides described to date appear to signal through GPCRs. Regardless of receptor type, ligands must be removed from the synapse following receptor activation before further signals can be transmitted, in a process known as signal termination. Peptidergic- and cholinergic-mediated signals are terminated by enzymes that destroy the transmitter within the synapse. All remaining classical transmitters are removed from the synapse by specific transporter pumps, which actively transport transmitters back into neurons or glial cells for metabolic breakdown or reuse.
Experiments performed from the 1960s onward demonstrated the presence in flatworms of a complement of now well-established classical/small-molecule neurotransmitters, including acetylcholine, dopamine, γ-aminobutyric acid, glutamate, histamine, noradrenaline, octopamine, and serotonin, all of which have been directly or indirectly detected in several flatworm species and/or have been localized to neuronal tissue (Ribeiro et al., 2005). Many also have demonstrable activity when added to whole flatworms or neuromuscular preparations in vitro. Although no existing drugs target classical neurotransmission in flatworms, the potential of this system as a source of therapeutic targets is highlighted by the many antinematode anthelmintics that target classical transmitter receptors. Dichlorvos, levamisole, morantel, piperazine, pyrantel, macrocyclic lactones, paraherquamide, and the amino acetonitrile derivatives (AADs) all act upon receptors or enzymes associated with classical neurotransmission (McVeigh et al., 2012).
Acetylcholine (ACh) is the primary excitatory transmitter at neuromuscular junctions of vertebrates and most invertebrates. It has species-specific excitatory or inhibitory neuromuscular impacts in flatworms, triggering muscle contraction in planarian preparations (Moneypenny et al., 2001; Blair & Anderson, 2009), while inhibiting myoactivity in trematode and cestode assays (Pax et al., 1984; Sukhdeo et al., 1984, 1986; Thompson et al., 1986; McKay et al., 1989; Day et al., 1996; Maule et al., 2009). In addition to its core neuromuscular role, there is also evidence that ACh is involved in nutrient acquisition at the host-parasite interface, since nicotinic ACh receptors (nAChRs) and acetylcholinesterase (AChE) are concentrated on the tegumental surface of adult male schistosomes. These cholinergic signaling elements are linked to schistosome glucose transporters, enabling glucose uptake from host blood, triggered by typical circulating concentrations of ACh. Glucose transport is ablated by both nAChR antagonists and AChE inhibitors (Camacho & Agnew 1995). This highlights an additional opportunity for anticholinergic compounds to target schistosomes; by targeting these surface-exposed elements, it may be possible to use preexisting pharmacology to interfere with parasite energy metabolism or glycolysis.
Biosynthesis of ACh from choline and acetyl coenzyme A is catalyzed by choline acetyltransferase (ChAT) (Fig. 1), with signaling terminated following receptor activation by the hydrolytic action of AChE. AChE activity can be detected histochemically, providing a convenient marker for cholinergic synapses—indeed, cholinesterase histochemistry provided the first evidence of ACh in flatworms (Graft & Read, 1967; Halton, 1967). ChAT, ACh's single biosynthetic enzyme, has been poorly studied in flatworms, with only a single publication reporting cloning and functional analysis of this protein from the planarian Dugesia japonica (Nishimura et al., 2010). Nishimura et al. (2010) employed RNA interference (RNAi) to silence the DjChAT gene, yielding an inhibited motility phenotype that phenocopied the impacts of nicotinic ACh receptor antagonist drugs. This effect is consistent with DjChAT's involvement in the synthesis of the excitatory ACh.
ACh is a ligand at both nicotinic (LGIC) and muscarinic (GPCR) receptors. Muscarinic ACh receptor (mAChR)-like sequences have been identified in cestode, planarian, and trematode genomes (Zamanian et al., 2011; Tsai et al., 2013; Campos et al., 2014). One mAChR (also termed GAR: G protein–coupled acetylcholine receptor) has been functionally expressed and characterized from S. mansoni (MacDonald et al., 2015) (Fig. 2). SmGAR expressed in yeast showed basal constitutive activity, with further activation triggered by the addition of ACh, but not tyramine, histamine, or glutamate, suggesting that SmGAR is a specific ACh receptor. RNAi analysis indicated that SmGAR has a role in neuromuscular coordination, since silencing the receptor inhibited the motility of larval S. mansoni (MacDonald et al., 2015) (Fig. 2).
Figure 2 Deorphanized flatworm receptors. Fourteen flatworm-derived receptors have been deorphanized in heterologous expression systems or by RNA interference (RNAi) (GtSER1 and GtNPR1 only). All are activated by classical transmitters except GtNPR1, which is a neuropeptide receptor. The ATP-gated channels (SchP2X and SmP2X), an acetylcholine-gated chloride channel (SmACC), and the glutamate gated chloride channels (SmGluCl) are ligand-gated ion channels; all of the remaining receptors are seven-transmembrane G protein–coupled receptors (GPCRs). Receptors are colored according to activity, where red represents a receptor with inhibitory function, green indicates excitatory function, and gray indicates unknown function. Heterologous expression systems, signaling pathways, function, and RNAi phenotypes are indicated where known. References: SchP2X (Agboh et al., 2004); SmP2X (Raouf et al., 2005); SmACC1 (MacDonald et al., 2014); SmGAR (MacDonald et al., 2015); SmD2 (Taman & Ribeiro, 2009); SmGPR3 (El-Shehabi et al., 2012); SmGluCl-1,-2,-3 (Dufour et al., 2013); SmGluR (El-Shehabi et al., 2009); SmGPCR (Fadi F Hamdan et al., 2002); SmGPR2 (El-Shehabi & Ribeiro, 2010); Sm5HTR (Patocka et al., 2014); Dj5HT7 (Nishimura et al., 2009); GtSER1 (Zamanian et al., 2012); and GtNPR1 (Omar et al., 2007; Zamanian et al., 2012).
Vertebrate nAChRs are invariably selective for cations, triggering membrane depolarization and cellular excitation when activated. The classic example of ACh-nAChR signaling is in the contraction of skeletal muscle at the vertebrate neuromuscular junction. Three nicotinic ACh receptor (nAChR) subunits have been cloned and localized in S. haematobium, one of which (ShAR2beta) was considered divergent based on the amino acids lining the ion pore (Bentley et al., 2004, 2007; MacDonald et al., 2014). ShAR1alpha and ShAR2beta are expressed in the S. haematobium tegument and musculature, respectively, but neither subunit formed functional channels, either alone, in combination, or coupled with a vertebrate alpha7 subunit, in Xenopus oocytes (Bentley et al., 2004). Given the availability of extensive nicotinic pharmacology that could potentially delineate parasite nAChRs from those of the host, the inability to functionally express these schistosome channels could be considered a missed opportunity for drug discovery. However, the presence of at least 12 nicotinic subunits in the S. mansoni genome (including homologues of the three S. haematobium subunits described earlier) (Berriman et al., 2009) could stimulate further efforts in this regard.
Invertebrates exhibit some interesting diversification of the ACh-gated ion channel complement; in addition to cation-selective nAChRs, they also possess anion (Cl–)-selective ACh-gated channels (ACCs). By transmitting negative charge, ACCs trigger cellular hyperpolarization and are therefore associated with inhibitory signaling. In mollusks and nematodes (Putrenko et al., 2004; van Nierop, 2005; Beech et al., 2013) ACC subunits are defined by the presence of a Pro-Ala anion-selective motif in the M2 domain that lines the channel pore. MacDonald et al. (2014) demonstrated the presence of this motif in five S. mansoni nAChR subunits. The anion-selective motif is also found in ShAR2 beta, the previously identified divergent nAChR subunit from S. haematobium (Bentley et al., 2007). Silencing ACC subunits in schistosomula yielded a hyperactive phenotype, consistent with the abolition of inhibitory signaling (Fig. 2). Heterologous expression of one subunit, SmACC-1, yielded a functional homomeric channel in HEK-293 cells that mediated anion influx upon exposure to nicotine and other cholinergic agonists. This response was inhibited by the nicotinic antagonist, D-tubocurarine, but not by other classical nicotinic antagonists (mecamylamine and atropine) (MacDonald et al., 2014). The tissue expression patterns of SmACC subunits have not yet been described, making it difficult to ascertain whether the motility impacts of SmACC RNAi are the result of silencing neuron- or muscle-expressed receptors. Nevertheless, these data suggest that ACC channel subunits may be responsible for at least some of the myoinhibitory effects of ACh on schistosomes (Day et al., 1996). Comparative studies on the presence and/or relative distribution and function of ACC in the planarian NS would be informative in the context of ACh's myoexcitatory actions in these turbellarians.
ACh signaling is terminated by the hydrolytic action of AChE present within cholinergic synapses. The potential of AChE as a drug target is highlighted by the success of organophosphate pesticides, which function as AChE inhibitors. Although now banned due to toxicity concerns, these compounds were extremely effective weapons against arthropods, nematodes, and other agricultural pest species, in which they caused muscular paralysis. AChEs from three Schistosoma spp. have been functionally expressed (Bentley et al., 2003, 2005). Importantly, these show structural differences from mammalian enzymes that might enable the development of compounds with selectivity between helminth and host AChE variants.
The catecholamine biosynthetic pathway is responsible for the production of dopamine, norepinephrine, and epinephrine. Catecholamines mediate a vast amount of CNS signaling in mammals, with roles in motor function, learning, and memory (Kobayashi, 2001). Dopamine has similarly central roles in invertebrates; in Drosophila, dopamine controls basal locomotion, as well as more complex functions in circadian rhythm, learning, and courtship behavior (Yamamoto & Seto, 2014). Dopamine is also involved in the regulation of molluskan feeding behavior (Baxter & Byrne, 2006). There is limited evidence for the presence of norepinephrine and epinephrine in invertebrate phyla, with the phenolamines (tyramine and octopamine) considered their invertebrate equivalents (Xu et al., 2015). Nevertheless, there is some evidence for the presence of both dopamine and noradrenaline in flatworms. Both transmitters, and their biosynthetic activities, have been identified in cestodes, planaria, and trematodes (Ribeiro et al., 2005). Dopamine and norepinephrine have distinct neuromuscular effects when applied to different parasitic flatworm species, including circular muscle relaxation (S. mansoni; Hillman & Senft, 1973; Tomosky et al., 1974; Mellin et al., 1983; Pax et al., 1984), muscle contraction (D. merlangi; A. G. Maule et al., 2009), and dual excitatory/inhibitory effects (F. hepatica; Holmes & Fairweather, 1984; Davis & Stretton, 1995; Ribeiro et al., 2005).
Dopamine is synthesized in a two-step reaction from tyrosine (Fig. 1), which is first converted to L-Dopa by tyrosine hydroxylase; L-dopa is subsequently converted to dopamine by dopa decarboxylase. Full-length, enzymatically functional examples of tyrosine hydroxylase have been cloned from schistosomes (Hamdan & Ribeiro 1998; Hu et al., 2011). Dopa decarboxylase (also known as aromatic L-amino acid decarboxylase) has not yet been reported in any flatworm, but basic searches of available flatworm genomes support this enzyme's existence in S. mansoni (Fig. 1). Norepinephrine is synthesized from dopamine by dopamine β hydroxylase (DBH). DBH has currently only been described in Schmidtea mediterranea, where it was used as a marker for transcription factors necessary for regeneration of noradrenergic neurons (Scimone et al., 2014). Figure 1 shows that a DBH homologue also appears in the S. mansoni genome. In mammals, epinephrine is produced from norepinephrine by phenylethanolamine N-methyltransferase (PNMT; Fig. 1). Neither epinephrine, nor PNMT activity, has been reported in any flatworm.
Dopamine signaling is transduced through GPCRs. In mammals, these receptors are classified into two families (D1 and D2 receptors), which themselves encompass five subtypes (the D1 family contains D1 and D5 subtypes; the D2 family comprises D2, D3, and D4 subtypes). D1 and D2 families are distinguished by signal transduction mechanism, where D1 receptors stimulate cAMP production through G protein Gαs/olf subunits, while D2 receptors inhibit cAMP through Gαi/o (Beaulieu & Gainetdinov, 2011). In flatworms, two dopamine-activated GPCRs have been identified from S. mansoni (SmD2 and SmGPR3; Fig. 2). SmD2 clusters phylogenetically with D2 sequences, suggesting D2-like sequence identity, but lacks the typical pharmacology of mammalian D2 receptors when challenged with dopaminergic agonists and antagonists. Most notable is the effect of apomorphine, a mammalian dopaminergic antagonist which acts as an agonist at heterologously expressed SmD2 (Taman & Ribeiro, 2009). SmGPR3 is one representative of a schistosome-specific clade of biogenic amine GPCRs lacking homologues in other genera (El-Shehabi et al., 2012), and it was detected throughout the schistosome nervous system in a pattern similar to that of dopamine (El-Shehabi et al., 2012). When exposed to catecholamines, heterologously expressed SmGPR3 was most potently activated by dopamine. When challenged with classical biogenic amine antagonists, SmGPR3's profile did not resemble that of any known mammalian dopaminergic receptor. Reminiscent of apomorphine's impact on SmD2, spiperone, a mammalian D2 antagonist displayed agonism of SmGPR3.
Catecholamines are removed from the synapse by transporter pumps. A single dopamine transporter (DAT) has been cloned and functionally characterized from S. mansoni. Recombinant SmDAT showed dopamine transport activity, but interestingly, displayed pharmacology more akin to the human norepinephrine transporter because of its low affinity for known DAT inhibitors, including cocaine (Larsen et al., 2011). This pharmacological difference may permit selective targeting of this transporter for therapy.
γ-aminobutyric acid (GABA) is synthesized from glutamate by glutamate decarboxylase (GAD; Fig. 1). GABA itself has been identified in high performance liquid chromatography (HPLC) analyses from cestode, planarian, and trematode nervous systems (Ribeiro et al., 2005). GAD biosynthetic activity has been described in adult S. mansoni (Mendonça-Silva et al., 2004). GAD has also been described in the planarian, D. japonica (DjGAD) (Nishimura et al., 2008a), as a marker for GABAergic neurons, where expression of both DjGAD mRNA and protein were localized to neurons of the brain and pharynx. Nishimura et al. (2008a) also used RNAi to silence DjGAD, triggering a reduction in detectable GABA, alongside an aberrant photosensitivity phenotype. DjGAD-knockdown ablated the negative phototaxis seen in wild-type worms, although their locomotion remained normal. These data suggest that GABAergic transmission is associated with planarian photosensitivity, but not motility.
GABA operates through two types of receptor—GABAA (GABA-gated Cl– channels (Vogt, 2015) and GABAB (GABA-activated GPCRs; Brown et al., 2015). Neither of these receptor types has been cloned or characterized from any flatworm. Indeed, Dufour et al. (2013) stated that they found no evidence for schistosome GABA receptors during database searches that led to their discovery of the first S. mansoni glutamate-gated Cl– channels. Similarly, three GPCR discovery papers make no mention of GABAB receptors in the S. haematobium, S. mansoni, and S. mediterrannea genomes (Zamanian et al., 2011; Tsai et al., 2013; Campos et al., 2014). There is some pharmacological evidence for GABAA receptors in schistosomes. Modulation of schistosome motility by picrotoxin, a GABAA receptor antagonist, implicates ionotropic GABA receptors in the control of schistosome motility (D. L. Mendonça-Silva et al., 2004), and there is evidence that benzodiazepine, a positive allosteric modulator of GABAA receptors, binds to schistosome extracts (Noël et al., 2007). These data suggest that GABAA receptors may be present in schistosomes, although their presence remains to be confirmed by cloning and functional characterization. No synaptic GABA transporters (GAT) have been formally identified in flatworms, but there is phylogenetic evidence for the presence of three GAT-like sequences in S. mansoni (Ribeiro & Patocka, 2013a).
Glutamate is synthesized from glutamine, by glutaminase (Fig. 1). This amino acid is ubiquitous throughout nature due to its obvious importance for protein synthesis, but glutamate is also an important neurotransmitter in both vertebrates and invertebrates. In vertebrates, glutamate is an excitatory transmitter that signals through both ionotropic glutamate receptors (iGluRs—LGICs) and metabotropic glutamate receptors (mGluRs—GPCRs). Invertebrates display additional functionality where glutamate may also trigger inhibitory signaling through glutamate-gated Cl– (GluCl) channels (Wolstenholme, 2012).
Ionotropic Glu receptors are composed of five subunit types: (i) AMPA (α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid); (ii) NMDA (N-methyl-D-aspartate); (iii) Kainate; and (iv) Delta subunits δ1 and δ2, termed "orphan subunits" because they have not yet been shown capable of forming a functional channel (Brockie, 2006). GluCl, AMPA, NMDA, Kainate, and Delta receptors exist in both vertebrates and invertebrates, (Brockie, 2006; Collingridge et al., 2009), while the GluCls are restricted to protostome invertebrates (Wolstenholme, 2012). Glutamate signaling is additionally transduced through mGluRs, of which there are eight subtypes (mGluR1-8) in mammalian systems (Niswender & Conn, 2010), and only one in Drosophila (Schoenfeld et al., 2013).
While none of the excitatory iGluR subunits have been cloned from flatworms, there is some pharmacological evidence for their presence. Kainate receptor pharmacology has been reported in schistosomes, radioactively labeled kainate binds to subcellular fractions of adult male S. mansoni, while addition of kainate to whole schistosomes induces a "corkscrew" motility phenotype (Mendonça-Silva et al., 2002). Similarly, both NMDA and AMPA induce seizure-like modulations of motility in planaria, similar to their impacts in mammalian models (Rawls et al., 2009). Four GluCl subunits have been cloned from S. mansoni (Dufour et al., 2013), three of which produced glutamate-gated Cl–-permeable channels when expressed in Xenopus oocytes (Fig. 2).
Several mGluRs are present in flatworm genomes—nine have been identified in S. mediterranea, three in Echinococcus multilocularis, and two each in S. mansoni and S. haematobium (Zamanian et al., 2011; Tsai et al., 2013; Campos et al., 2014). One of these S. mansoni receptors was cloned, functionally expressed, and localized to the central and peripheral nervous systems of adult and larval worms, and the adult female reproductive tract (Taman & Ribeiro, 2011b). This receptor was activated by glutamate, and it was unresponsive to GABA (Fig. 2). Importantly, the receptor's response profile to classical glutamatergic modulators suggests that it has quite different pharmacology from mammalian mGluRs, indicating its potential as a target for schistosome-specific drugs. An additional truncated mGluR has been reported in S. mansoni (SmGBP) (Taman & Ribeiro, 2011a), consisting of the extracellular N-terminal ligand binding domain, but lacking the heptahelical transmembrane domain that is characteristic of these receptors. SmGBP binds glutamate in radiolabeled glutamate competition assays. Expressed solely in the surface tubercles of adult male worms, Taman and Ribeiro (2011b) hypothesize a role in host-parasite signaling for SmGBP, although since it lacks the structural features required for G protein interaction, this truncated GluR must operate via another, potentially novel, signaling mechanism. Truncated GluRs were identified in other flatworms but have not been described in other metazoans (Taman & Ribeiro, 2011a). Metabotropic glutamate receptors are known to form functional dimers, where such interactions are essential for signaling (Pin & Acher, 2002; Kniazeff et al., 2004; Kammermeier & Yun, 2005). Immunoprecipitation experiments showed that SmGBP forms noncovalent dimers at neutral pH; Taman and Ribeiro hypothesized that SmGBP may regulate glutamatergic signaling by interacting/dimerizing with full-length mGluRs.
In cestode preparations, glutamate addition triggers a range of neuromuscular impacts, including contraction of muscle strips, increased electrical signaling in nerve tissue, and production of inositol triphosphate (IP3) second messengers (Ribeiro et al., 2005), the latter of which implicates the involvement of mGluRs in cestode glutamatergic signaling. The impact of glutamate on other flatworms is less compelling (Ribeiro et al., 2005). Aside from the aforementioned reports of kainate activity in schistosomes (Mendonça-Silva et al., 2002) and AMPA/NMDA impacts on planarian motility (Rawls et al., 2009), there is no strong evidence for the direct involvement of glutamate receptors in the motility of monogeneans, planarians, or trematodes. In the latter case, although glutamate does cause isolated schistosome muscle fibers to contract, this is thought to be an electrogenic, rather than receptor-mediated effect (Miller et al., 1996).
Little is known of flatworm synaptic glutamate transporters (GLTs), aside from a bioinformatics analysis that predicts a single GLT in the S. mansoni genome, of the solute carrier (SLC1) family (Ribeiro & Patocka 2013a).
Histamine biosynthesis is performed from histidine by histidine decarboxylase (Fig. 1). Although some flatworms are known to possess this enzyme and produce endogenous histamine, others are thought to obtain histamine from their host (El-Shehabi & Ribeiro, 2010). Histamine acts exclusively through GPCRs, which in humans comprise four families, H1–H4 (Stark, 2007). Exogenous applications of histamine have generally positive impacts on motility of live, intact cestodes and trematodes, and some H1 antagonists (H2, H3, and H4 blockers have no effects) have correspondingly inhibitory effects on motility (Ribeiro et al., 2005). Histamine lacks impacts on the motility of flatworm neuromuscular assay preparations (i.e., exposed neuromuscular assays in the form of muscle strips, trimmed worms, or dispersed muscle fibers). In line with these data, it has been suggested that rather than acting as an endogenous signaling molecule, histamine's stimulation of parasite motility may in fact enable the parasite to respond to the presence of host histamine, levels of which increase following infection with schistosomes (Ribeiro et al., 2005). Given histamine's role in host immune signaling, such ability would be advantageous, particularly for blood-dwelling schistosomes. Consistent with this hypothesis, one of the two known schistosome histamine receptors (SmGPCR, Fig. 2; Hamdan et al., 2002) localizes to the tegument of schistosomula, adult male and intramolluskan sporocyst forms of S. mansoni. The second S. mansoni histamine receptor (SmGPR2, Fig. 2; El-Shehabi & Ribeiro, 2010) localizes to the nervous system, suggesting an endogenous neuromuscular role. Both of these GPCRs are activated by histamine in heterologous expression systems (Fig. 2). Profiling SmGPR2 with classical histamine antagonists (El-Shehabi & Ribeiro, 2010) reveals distinct pharmacology from mammalian histamine receptors, hinting at the ability to specifically target this receptor.
Synaptic inactivation/removal of histamine occurs by an unknown mechanism, since no known biogenic amine transporter has been shown to exhibit histamine transport activity. Histamine reuptake may instead be performed by unselective organic cation transporters of unknown identity (Ogasawara et al., 2006; Ribeiro & Patocka, 2013b).
Nitric oxide (NO), a gaseous signaling molecule in vertebrates (Ignarro, 1990), has been indirectly identified in flatworms based on the presence of its biosynthetic enzyme, nitric oxide synthase (NOS). NOS synthesizes NO from L-arginine (Fig. 1). NO signals via activation of soluble guanylate cyclase, which catalyzes the production of a cyclic GMP (cGMP) second messenger that primarily activates protein kinase G. In vertebrates, this signaling axis performs neurotransmission, as well as regulating aspects of cardiovascular and renal function (Bian & Murad, 2014). NOS, as visualized by histochemical staining of nicotinamide adenine dinucleotide phosphate (NADPH) diaphorase activity (Hope et al., 1991), localizes to neurons in both cestodes and trematodes (Ribeiro et al., 2005). Localization of NOS in adult schistosomes additionally identified NOS immunoreactivity in the surface-exposed gut and tegument (Kohn et al., 2001). Expression of NOS at the parasite surface may suggest a role for NO signaling at the host-parasite interface (Da'dara & Skelly, 2011). Host-derived, NO-mediated signals may be received by schistosomes, given that exogeous NO can affect schistosome transcriptional profiles (Messerli et al., 2006). At present there are no data regarding NO's function in the flatworm NS or in the parasite's ability to influence the host environment.
Phenolaminergic transmitters comprise tyramine and octopamine. Both of these transmitters are synthesized from a single sequential pathway (Fig. 1) in which tyramine is produced from tyrosine by tyrosine decarboxylase, which itself is subsequently converted to octopamine by tyramine ß-hydroxylase (TBH). Tyramine and octopamine are considered the invertebrate equivalent of the vertebrate adrenergic transmitters (epinephrine and norepinephrine) (Xu et al., 2015).
Tyramine is an established neurotransmitter in insects (Roeder et al., 2003; Roeder, 2005). Its existence in flatworms is implied by the presence of tyrosine decarboxylase activity in cestodes (Ribeiro & Webb, 1983) (Fig. 1). However, since tyramine is also an essential intermediate in octopamine biosynthesis, and its neuro-/myoactivity in any flatworm assay has yet to be demonstrated, its status as an endogenous flatworm neurotransmitter remains uncertain. The occurrence of octopamine in flatworms is similarly implied by the presence of TBH activity in Hymenolepis diminuta extracts (Ribeiro & Webb, 1983). TBH has also been cloned from the planarian Dugesia, where knockdown reduced the amount of HPLC-detectable octopamine in tissue extracts (Nishimura et al., 2008b). Nishimura et al. are the only authors to have performed RNAi on any aspect of the phenolaminergic signaling pathway, and it is notable that they reported no aberrant behavior or motility phenotypes associated with TBH ablation. This may suggest that octopamine, in planarians at least, is not involved in neuromuscular function, or at most has a subtle role that does not obviously influence gross motility or behavior. Octopamine and tyramine both act via GPCRs in invertebrates (Farooqui, 2007; Lange, 2009), although homologues of these receptors have not yet been reported in flatworms. Neither have any synaptic phenolamine transporters been identified in flatworms, although one of the S. mansoni biogenic amine transporters has been tentatively annotated as an octopamine transporter (Caveney et al., 2006; Ribeiro & Patocka, 2013b).
Serotonin (5-hydroxytryptamine; 5HT) is the best studied (in terms of publication volume) of all flatworm classical neurotransmitters. It has been identified in all four platyhelminth classes and appears abundant throughout the entire nervous system (Ribeiro et al., 2005). Biosynthesis of 5HT involves a two-step reaction from tryptophan (Fig. 1), which is first converted to 5-hydroxytryptophan by tryptophan hydroxylase (TPH), from which 5HT is produced by L-aromatic amino acid decarboxylase (L-AADC). There is biochemical evidence for both of these enzymes throughout phylum Platyhelminthes (Ribeiro et al., 2005). An S. mansoni homologue of TPH (SmTPH) has been cloned and functionally characterized in vitro (Hamdan & Ribeiro, 1999). Although the human and schistosome enzymes are biochemically similar, the mammalian enzyme is difficult to study in vitro due to extreme instability, whereas SmTPH is stable at 37°C. Consistent with 5HT's constitutive expression, SmTPH transcripts are detectable in all life stages (Hamdan & Ribeiro, 1999; Boyle et al., 2003). L-AADC (also known as Dopa decarboxylase, where it is also involved in dopamine synthesis—see earlier) has not yet been reported in any flatworm.
5HT is considered the flatworm primary excitatory neurotransmitter. In in vitro assays, exogenously applied 5HT stimulates motility of whole worms and causes muscle contraction in trimmed worms and muscle strips (Ribeiro et al., 2005). On dispersed muscle fibers, 5HT has distinct impacts between schistosome- and planarian-derived preparations. Muscle fibers from the planarian Procerodes littoralis contract when directly exposed to 5HT, in a concentration-dependent manner, while preincubation in serotonin blunts the contractile responses of other myoactive agents, such as FMRFamide-related neuropeptides (Moneypenny et al., 2001). In contrast, serotonin does not cause S. mansoni fibers to contract, but it is a prerequisite for depolarization-induced contraction of schistosome-derived fibers (Day et al., 1994). Although technical differences between these two assays (that employ distinct methods in liberating cells from each species) cannot be ruled out as an explanation for the apparent disparity, these data could indicate the existence of species-specific differences in flatworm serotoninergic neuromuscular physiology. There is also ample evidence for serotoninergic signaling having an important role in platyhelminth developmental processes. As well as evidence linking 5HT and 5HT receptors with planarian cellular regeneration (see Ribeiro et al., 2005, for review), serotonin similarly stimulates the differentiation of protoscoleces toward metacestodes in Echinococcus spp. cestodes (Camicia et al., 2013). The lack of a body cavity and a circulatory system in acoelomate flatworms designates the nervous system as the primary carrier of developmental cues. As such, neurotransmitters could well have developmental signaling roles in addition to those responsible for controlling neuromuscular and sensory functions. We would anticipate similar roles in the control of regeneration and cellular renewal for other transmitters as well as 5HT, although this hypothesis has not been adequately tested.
Mammalian 5HT receptors comprise seven major types (5HT1–7); one (5HT3) is an LGIC, whereas the remainder are GPCRs (D. E. Nichols & Nichols, 2008). The GPCR subtypes are distinguished by signal transduction pathway, where 5HT1 and 5HT5 receptors signal through Gαi/o, by inhibition of cAMP, 5HT2 stimulates IP3 synthesis through Gαq/11, while 5HT4, 6, and 7 stimulate cAMP production via Gαs. Invertebrate 5HT receptor families appear somewhat less complex; C. elegans, D. melanogaster, and Apis mellifera all possess 5HT1, 2, and 7 orthologues (Dernovici et al., 2007; Blenau & Thamm, 2011). C. elegans appears unique in also employing a 5HT-gated Cl– channel known as MOD-1 (Ranganathan et al., 2000). Three 5HT-activated GPCRs have been functionally described in flatworms (Fig. 2). The first, from D. japonica, was activated by 5HT when functionally expressed in Xenopus oocytes. Dj5HT7 (named for its homology to mammalian 5HT7), signaled through a Gαs-compatible chimeric G protein and did not respond to other neurotransmitters aside from 5HT (Nishimura et al., 2009). A second planarian (Dugesia tigrina) 5HT receptor was identified using a novel, RNAi-guided approach to deorphanization (Zamanian et al., 2012). By performing RNAi on individual receptors alongside a second messenger assay as a post-RNAi phenotypic readout, Zamanian et al. demonstrated that DtSER-1 knockdown reduced the levels of 5HT-stimulated cAMP generated by tissue homogenates. This novel methodology illustrated that DtSER-1 was activated by serotonin, without the need to clone and functionally express the receptor in a heterologous system. The latter, more traditional method was used by Patocka et al. (2014) to deorphanize a 5HT7-like receptor from S. mansoni (Sm5HTR). Sm5HTR is expressed throughout the CNS and PNS, and it is involved in parasite motility. The receptor specifically appears involved in transducing excitatory neuromuscular signals, since Sm5HTR RNAi drastically reduces motility in both larval and adult schistosomes.
Serotonin signal termination occurs by physical removal of 5HT from the synapse, a process performed by serotonin-specific reuptake transporter (SERT) pumps. These pumps are pharmacologically important targets for commonly prescribed antidepressant drugs, including specific serotonin reuptake inhibitors (SSRIs), such as fluoxetine and citalopram, and tricyclic antidepressants, such as imipramine (Gether et al., 2006). The druggability of these transporters, alongside their involvement in demonstrably important neuromuscular functions, highlights their obvious appeal as anthelmintic targets. One SERT has been identified and functionally characterized in S. mansoni (Patocka & Ribeiro, 2007, 2013; Fontana et al., 2009; Ribeiro & Patocka, 2013b). SmSERT is represented by two isoforms. While sensitive to many of the drugs that act on human SERT, the relative potencies of these drugs are quite different on helminth versus host transporters. This indicates the presence of distinct pharmacology that could be exploited to develop parasite-specific SERT-targeting drugs.
Since the 1950s there have been suggestions that ATP was released by neurons following electrical stimulation (Mutafova-Yambolieva & Durnin, 2014). There is now an appreciation that purinergic signaling is involved in many physiological processes, including neurotransmission. Within the mammalian brain, purinergic signaling is responsible for neuromodulation, synaptic plasticity, and even fast synaptic transmission (Burnstock, 2013), processes that are mediated by two types of purinergic receptor—P2X trimeric ionotropic receptors (Wang & Yu, 2016) and P2Y GPCRs (von Kügelgen & Hoffmann, 2015). Following receptor interaction, ATP hydrolysis and signal termination is performed by one of eleven ecto-nucleotidases (Zimmermann & Braun, 1999; Williams, 2002). Limited research attention has been paid to the purinergic signaling components of flatworms. The only evidence for these pathways comes from two reported P2X receptors in S. mansoni (Agboh et al., 2004; Raouf et al., 2005). Since the expression profiles of these receptors remain unknown, their functions remain similarly cryptic. Aside from neurotransmission, schistosome P2X receptors could conceivably be surface expressed and responsible for detecting host ATP in some aspect of host-parasite communication. Clearly, further research is required to understand the importance, if any, of purinergic neurotransmission in flatworms.
Classical neurotransmitters are clearly involved in important aspects of platyhelminth neuromuscular function, and their receptors and synaptic transporters represent validated anthelmintic targets, as evidenced by the RNAi and pharmacological phenotypes detailed earlier. Arguably as significant in terms of function is the peptidergic arm of the flatworm nervous system. Immunocytochemical studies demonstrate that the extent of neuropeptide immunoreactivity outstrips that of classical transmitters in flatworm neuronal elements (Halton & Maule, 2004), and all neuropeptides tested on flatworm bioassays have invariably been biologically active (exhibiting either myoactivity or stimulation of signaling activity). Allied to clear differences in peptide signaling and function between parasites and host organisms, these factors argue in favor of flatworm peptidergic signaling as a potential source of targets for new anthelmintic drugs (Mousley et al., 2005; McVeigh et al., 2012). Although well characterized in terms of peptide distribution, physiology, and pharmacology, the flatworm neuropeptidergic nervous system has only in recent years benefited from improved sequence datasets, which have allowed the first characterizations of the genes encoding putative neuropeptides, peptide receptors, and associated signal transduction proteins. Here, we will summarize current understanding of the diversity and function of flatworm neuropeptides and their biosynthetic and signaling elements.
Helminth neuropeptide synthesis has been extensively reviewed elsewhere, and the reader is referred to these reviews for graphical representations of the helminth neuropeptide biosynthetic pathway (McVeigh et al., 2005a, 2005b; Husson & Schoofs, 2007a, 2007b; Li & Kim, 2008; Geary, 2010).
Neuropeptides are transcribed and translated from their encoding genes on precursor proteins, known as prepropeptides, that may incorporate single or multiple copies of peptides, with the latter often bearing related motif sequences. Prepropeptides display an N-terminal secretory signal peptide sequence that targets the protein into the cellular secretory pathway. During ribosomal translation, this sequence is the first part of the nascent protein to emerge from the ribosome, enabling its binding to a signal recognition particle (SRP). The SRP translocates the growing protein and its ribosome to the ER, where the protein continues to be synthesized through a translocon (a channel in the ER membrane) into the ER lumen. Once in the lumen, the signal peptide sequence itself is cleaved from the preproprotein. Subsequent biosynthesis occurs within the ER and Golgi, and it involves excision of peptides from the precursor by prohormone convertase enzymes, and the removal of remaining dibasic cleavage markers by carboxypeptidases. The resulting peptides are then available for posttranslational modifications, the most common of which is C-terminal amidation, which converts a C-terminal Glycine amino acid (found at the C-terminus of many encoded/immature neuropeptides) to amide (NH2). Amidation in flatworms is performed by the sequential actions of two enzymes, peptidylglycine α-hydroxylating monooxygenase (PHM) and peptidyl α-hydroxyglycine α-amidating lyase (PAL). In mammals, both of these functions are performed by a single bifunctional enzyme, peptidylglycine α-amidating monooxygenase (PAM). Amidated peptides are considered mature. They are stored in presynaptic secretory vesicles, from which they are exocytosed into the synapse when required.
All flatworm neuropeptides are synthesized essentially by the mechanisms described earlier, but they display important differences in structure and function derived from primary amino acid sequence and the presence/absence of posttranslational modifications. Recent years have seen improved understanding of the sequence diversity of flatworm neuropeptide genes, thanks largely to improved genomic sequence resources. Prior to the genomic era, neuropeptide structure and sequence data were derived from HPLC-based isolation and peptide sequencing of peaks/fractions with immunoreactivity to known peptides from other species (Day & Maule, 1999). These methods yielded seven flatworm peptides—five FMRFamide-like peptides (FLPs) (Maule et al., 1993, 1994; Johnston et al., 1995, 1996) and two neuropeptide Fs (NPFs) (Maule et al., 1991; Curry et al., 1992). These isolations are particularly impressive given the technical difficulties associated with detecting peptides that are generally of low abundance from, as is the case with parasite-derived peptides, biological samples that are difficult to obtain in large quantities. For example, the extraction of GNFFRFamide from Moniezia expansa required one kilogram of tissue (Maule et al., 1993). Given these technical limitations, it is perhaps understandable why no peptides have been biochemically isolated from any trematode species. Subsequent work employed molecular techniques to clone the genes encoding some of these peptides in cestodes, trematodes, and planaria (Mair et al., 2000; Dougan et al., 2002; Humphries et al., 2004), but not until the arrival of the genomic era did we appreciate the full complexity of the flatworm neuropeptidergic system. Table 1 illustrates our current knowledge of neuropeptide precursor genes in phylum Platyhelminthes, based on published data derived from genome or transcriptome datasets. Sixty-four neuropeptide-encoding genes are known in the phylum, although these data are derived from only 12 species, and the distribution of homologues between species remains uncertain. It is unclear whether gaps in the interspecies distributions of npp homologues indicate a species-specific component of neuropeptide expression, or simply the incomplete nature of many of these datasets. S. mediterrannea represents the current most complete dataset, as reflected by its 51 precursors and 142 validated peptides (Collins et al., 2010). MS-validated peptides from this study were also queried against the genomes of S. japonicum and S. mansoni, allowing Collins et al. to identify an additional eight schistosome prepropeptide genes, including the YIRFamide encoding npp-23. YIRFamide was originally isolated from Bdelloura candida, a marine turbellarian, and is one of the most commonly tested peptides in flatworm bioassays (see later). Native Schmidtea peptides have not yet been searched against other flatworm datasets, so their conservation across the phylum remains unknown.
Notes. Predicted peptide encoding genes are shown, alongside the peptide family encoded by each gene, as described by (a) McVeigh et al., 2009, (b) Berriman et al., 2009, (c) Collins et al., 2010, and (d) Tsai et al., 2013. Representation of each gene in genome or transcriptome datasets is identified by shading. Where more than one copy of a gene is present (i.e., two or more genes encoding essentially identical peptides), the number of copies is denoted by a number. Asterisks identify genes from which predicted peptides have been validated by mass spectrometry (Collins et al., 2010).
cpp, cerebral peptide prohormone; grh, gonadotropin releasing hormone-like; ilp, insulin-like peptide; mpl, myomodulin prohormone-like; npp, neuropeptide precursor; npy, neuropeptide Y superfamily; ppl, pyrokinin prohormone like; ppp, pedal peptide prohormone like; spp, secreted peptide prohormone.
The flatworm neuropeptide dataset contains homologues of recognized invertebrate peptide families, including the FLP and NPF/NPY families, as well as APGWamide, buccalin, and myomodulin. S. mansoni also encodes peptides very similar to vertebrate neuropeptide FF (McVeigh et al., 2009). Schmidtea possesses additional homologues of cerebral peptide, gonadotropin-releasing hormone, insulin-like peptide, pyrokinin, and pedal peptide precursors (Collins et al., 2010). Based on the varied roles of their homologues in other invertebrates, it seems reasonable to posit that flatworm peptides fulfil a broad range of functions in addition to the more commonly ascribed neuromuscular roles of FLPs. Notably, with the exception of FLPs and NPFs, most of these peptide families have not yet been tested in flatworm neuromuscular bioassays.
Silencing components of the neuropeptide biosynthetic pathway illustrates the importance of peptidergic signaling for flatworms. RNAi of a PC2-like prohormone convertase in S. mediterrannea correlates with globally reduced levels of properly processed peptides in MS-analyzed tissue extracts. This intervention interferes with normal motility/behavior, regeneration, and reproductive development (Reddien et al., 2005; Collins et al., 2010). Similar phenotypes were reported following knockdown of a PC2 chaperone protein (7B2) and regulator of G protein signaling (RGS) proteins (Reddien et al., 2005), suggesting that neuropeptide biosynthesis and function are generally important for normal helminth behavior. PC2 knockdown has also been used to demonstrate a link between neuropeptide signaling and pharynx extension in planarians (Shimoyama, Inoue, & Kashima, 2016).
FLPs (referred to in early literature as FMRFamide-related peptides, FaRPs), represent the largest known family of bioactive peptides in nature (McVeigh et al., 2006). Originally named based on sequence similarity with the eponymous FMRFamide, a cardioactive tetrapeptide in mollusks, FLPs are defined as short, bioactive amidated peptides with a C-terminal RFamide motif (McVeigh et al., 2009). Such peptides are now recognized as common throughout the protostomian invertebrate lineage. In deuterostomes the family has diversified to encompass several subfamilies bearing C-terminal RFamide motifs (e.g., gonadotropin inhibitory hormone, neuropeptide FF, prolactin-releasing peptide) (Elphick & Mirabeau, 2014). Flatworms possess at least seven FLP-encoding gene homologues (Table 1).
FLPs trigger a consistent excitatory/contractile response in whole-worm assays, muscle strip preparations, and dispersed muscle fibers from several species, with qualitatively similar excitatory responses also triggered by FLPs of nematode or arthropod origin (Mousley et al., 2004). The universally excitatory impact of FLPs suggests the existence of a monofunctional FLP receptor-signaling pathway in flatworms, data that are particularly intriguing given the multifunctional motility impacts of FLP families in other invertebrate genera (Mercier et al., 2003; R. Nichols, 2003; López-Vera et al., 2008). An important advance was the development of a muscle cell dispersal assay that enabled peptides to be tested on functional muscle cells that had been enzymatically digested and separated from other tissue (Blair et al., 1991). Significantly, this assay allowed experimentation in the absence of neuronal tissue; in the case of whole-worm and muscle strip assays, the relative impacts of test compounds on nerve and muscle cannot be discerned. Dispersed cell experiments illustrated the presence of a FLP-activated receptor on flatworm muscle tissue, implicating FLPs in the direct control of flatworm neuromuscular function (Day et al., 1994, 1997; Moneypenny et al., 2001).
FLPs in other invertebrates exert their functions primarily through GPCRs (Nichols, 2003; Claeys et al., 2005; Peymen et al., 2014; Zatylny-Gaudin & Favrel, 2014). A notable exception is the FMRFamide-activated Na+ channel (FaNaC), through which FMRFamide mediates fast neuronal transmission in gastropod mollusks and cnidarians (Lingueglia et al., 1995; Golubovic et al., 2007). There is also some electrophysiology evidence for a FLP-gated Cl– channel in nematodes (Maule et al., 1995; Holden-Dye et al., 1997; Purcell et al., 2002). There is currently no evidence for peptide-gated ion channels in flatworms.
Only one FLP-activated flatworm GPCR has been directly identified, from the planarian D. tigrina (GtNPR1; Omar et al., 2007; Zamanian et al., 2012). In a heterologous expression system, this receptor responded most potently to GYIRFamide (Fig. 2). Although also activated by peptides encoded by the nematode flp-1 gene, GtNPR1 did not respond to flatworm NPFs. A subsequent RNAi-based deorphanization study confirmed GYIRFamide's activation of GtNPR1 and showed that endogenous GtNPR1 signaled through a Gαi/o pathway (Zamanian et al., 2012; Fig. 2). Although these studies are promising demonstrations of the feasibility of flatworm peptide receptor deorphanization, further work is required before GtNPR1 or its parasite homologues can be considered as candidate anthelmintic drug targets. Notably, Zamanian et al. (2012) reported no phenotypic consequences of silencing GtNPR1, although it is unclear whether detailed phenotypic screens were performed in this study. In any case, the absence of obviously aberrant motility phenotypes suggests that GtNPR1 is probably not a muscle-based FLP receptor; GtNPR1 currently lacks tissue expression data that would either support or refute this hypothesis. Further studies are required to clarify both of these points. In terms of localization, we also lack any evidence that GYIRFamide and GtNPR1 are expressed in a manner that would allow their interaction in vivo. One major advantage of GtNPR1 as a target for compound screens is its clear amenability to heterologous expression in a high throughput screen (HTS)-compatible system. The argument for committing GtNPR1 to HTS of pharmaceutical compound libraries could be solidified by addressing the knowledge gaps pointed out earlier. These data would help ascertain GtNPR1's candidacy by contributing to a "validation portfolio" of relevant biological data that would permit evaluation of the potential of this parasite receptor as a drug target candidate.
In addition to GtNPR1, which signals through Gαi/o and adenylate cyclase (Zamanian et al., 2012), there is biochemical evidence for additional FLP-activated signaling pathways in trematodes. In F. hepatica, GYIRFamide activates a phospholipase C (PLC)/protein kinase C (PKC)–coupled receptor pathway in tissue homogenates (Graham et al., 2000). Given the crude nature of the whole-worm extracts used in these assays, it is impossible to ascertain from which tissues these receptor signals originated. Although employing very different methods, a comparison of Graham et al. (2000)'s results with those of Zamanian et al. (2012), discussed earlier, suggests that GYIRFamide operates through distinct GPCR signaling axes in trematodes (Gαi/o-adenylate cyclase) and planarians (Gαq-PLC). If such interspecies distinctions hold true for other receptors, we may find that planarians are of limited utility as models for parasitic forms.
Novozhilova et al. (2010) demonstrated that YIRFamide's contractile effect on S. mansoni muscle fibers was mediated through a PKC-dependent pathway, triggering cytosolic Ca2+ influx through sarcolemmal voltage-operated Ca2+ channels (VOCCs).
There is appreciable evidence of a central role for FLPergic signaling in flatworm neuromuscular function, and these data frame flatworm FLP signaling as a source of potential therapeutic targets. We still lack many of the core tools required to exploit neuropeptide receptors and associated proteins for drug discovery, most notably the absence of any deorphanized parasite FLP receptors.
NPFs have been isolated biochemically from two flatworm species, the cestode Moniezia expansa and the turbellarian Arthurdendyus triangulatus (Maule et al., 1991; Curry et al., 1992). Attempts to isolate NPF from schistosomes were unsuccessful even when using 10,000 adult S. mansoni (Humphries et al., 2004). These isolations were stimulated by studies reporting immunoreactivity to members of the vertebrate neuropeptide Y superfamily in flatworm nervous systems, particularly to pancreatic polypeptide (PP). Chromatographic experiments showed that the single PP immunoreactive peaks in extracts from Diclidophora merlangi, M. expansa, and S. mansoni coeluted with other NPF/NPY peptides, suggesting that these peaks contained peptides with NPY superfamily-like sizes and hydrophobicities, and implying the presence of NPF-like peptides in these species (McVeigh et al., 2005b). Extraction of cestode and turbellarian NPFs, and subsequent cloning of their encoding genes, provided the first evidence of flatworm NPF structure. Flatworm NPFs are 29–47 amino acids in length, with conserved motifs consisting of a C-terminal GRPRFamide, with conserved tyrosines at positions 10 and 17 upstream of the C-terminus. Classically, NPFs (named for their C-terminal Phe) were considered the invertebrate version of NPY (named for C-terminal Tyr), which itself was considered restricted to vertebrates. Genomic data have now revealed that both NPF and NPY peptides can be present in individual flatworm species. Whether there are differences in function or signaling between these peptides remains unknown. Genomic structure provides further evidence for the relatedness of NPF/NPY, since almost all NPF and NPY encoding genes described to date have a conserved phase 2 intron in the codon for their penultimate Arg residue (Larhammar et al., 1987; Mair et al., 2000; Dougan et al., 2002; McVeigh et al., 2009; Collins et al., 2010).
NPY superfamily peptides signal exclusively through GPCRs. Various NPY receptor subfamilies are recognized, including Y1, Y2, Y4, Y5, and Y6 (Larhammar & Salaneck, 2004), each of which varies in its occurrence and complexity between vertebrate species. Invertebrate NPF receptors are also GPCRs, but they are characterized by lower gene complexity—only one or two NPF receptors are present in insect and mollusk genomes (Nässel & Wegener, 2011; Vogel et al., 2013). NPF is involved with functions relating particularly to feeding/appetite, as well as learning, stress, and alcohol tolerance in insects and mollusks (Nässel & Wegener, 2011). While no flatworm NPF receptors have been cloned, available evidence suggests that NPFs trigger conserved NPY-like signaling pathways. NPY signals through Gαi/o by inhibiting the accumulation of cAMP (Motulsky & Michel, 1988; Aakerlund et al., 1990). Similarly in schistosome homogenates, NPF inhibits the accumulation of cAMP at concentrations as low as 10–11 M (Humphries et al., 2004).
Data on the functions of flatworm NPFs are similarly sparse. There is some evidence from S. mediterranea to implicate an NPF (Smed-npy-8), and its receptor, in reproductive function (Collins et al., 2010; Saberi et al., 2016). Knockdown of these genes led to phenotypes indicating a loss of sexual maturity, including regressed testes, cessation of sperm production, regression of copulatory organs, and in some cases, the disappearance of the gonopore. These physical phenotypes correlated with interrupted expression of developmental genes in affected tissues, suggesting that NPY-8 is essential for the normal maintenance of reproductive maturity that is required for planarian reproduction. NPF may also have a role in the control of cellular regeneration in planarians, given that exogenously applied NPF stimulates the mitotic activity of neoblasts, which translates to accelerated regeneration of the head, nervous system, and musculature (Kreshchenko et al., 2008).
Peptides with L/M/Iamide C-termini comprise the largest neuropeptide family in flatworms, with such peptides encoded by 21 of the 64 known neuropeptide genes (Berriman et al., 2009; McVeigh et al., 2009; Collins et al., 2010; Tsai et al., 2013). Some known peptides bear this motif, including molluskan buccalin (Aplysia: GMDSLAFSGGLamide; Cropper et al., 1988), myomodulin (Aplysia: PMSMLRLamide; Cropper et al., 1987), and arthropod pyrokinin (Leucophora: GTSPTPRLamide; Holman et al., 1986) neuropeptides. Buccalin, myomodulin, and pyrokinin are all myoexcitatory in their species of origin, so we would hypothesize that their flatworm homologues are similarly myoactive. Only one study has examined the bioactivity of a native flatworm member of this family. GFVRIamide, one of the peptides produced by the npp-1 gene, localizes to two neurons in the schistosome CNS and has inhibitory impacts on motility following application to whole-adult S. mansoni (McVeigh et al., 2011). Attempts to silence Sm-npp-1 were uninformative; although npp-1 transcripts were potently and specifically knocked down in schistosomulae, no aberrant motility phenotypes were noted in these worms. Arthropod type-A allatostatins also bear C-terminal L/Iamide signatures (characteristic motif: (Y/F)XFG(L/I)amide) and have myoactive effects in their host species. Allatostatins also trigger contractions of flatworm dispersed muscle fibers, in an example of interphyla functionality (Mousley et al., 2005).
Several additional smaller neuropeptide families have also been described from flatworms (McVeigh et al., 2009; Collins et al., 2010), for which limited data are available. These include the PWamides encoded by npp-5, which have been localized immunocytochemically to the S. mansoni PNS, including in innervation of muscle and reproductive organs (McVeigh et al., 2009). Although no somatostatin-like peptides have been identified, there is some evidence for the presence of somatostatin receptor-like GPCRs on the S. mansoni tegument (Chatterjee et al., 2005, 2007).
The neurobiology of parasitic flatworms appears to offer many opportunities for chemotherapeutic targeting in pathogenic helminths. The rapid development of genomics/transcriptomics, functional genomics, and heterologous expression methods during the last decade have enabled helminth parasitology to move from studying putative therapeutic targets based almost entirely on biochemistry, localization, and/or neuromuscular physiology toward the current paradigm that aims to apply systems biology methods to the identification, validation, and exploitation of potential chemotherapeutic targets. As detailed herein, these approaches have identified a cohort of heterologously expressed receptors, ion channels, and reuptake pumps, a handful of which have additionally been functionally validated by RNAi. These data, taken alongside improved knowledge of neuropeptides and biosynthetic enzyme complements, represent tangible progress in flatworm neurobiology during the past decade. Nevertheless, much remains to be done if this promise is to be translated into concrete progress with respect to developing new therapeutic options with which to treat helminth infections.
The receptors, transporters, and ion channels discussed earlier represent eminently druggable proteins, for which multiple classes of active compounds already exist. In addition to this appreciable pharmacopeia, it is particularly noteworthy that all of the proteins detailed earlier as successfully expressed in heterologous systems are described as "pharmacologically distinct" from their mammalian homologues. This implies considerable promise in the potential to target these proteins in a selective manner that would avoid nonspecific impacts on the host.


Wildflower Walks - Featuring Fiddleheads at Letchworth State Park!
Become more familiar with our many native wildflowers and the role of their surroundings, including the other plants and animals in their daily lives. Enjoy the fascinating variation in floral structure and folklore. Meet at Silver Lake Outlet Bridge, one mile north of Perry Exit on Main Park Road. 3 hours, 1 mile.


I gave some background about the struggles I am talking about today in yesterday's post. If you missed it, I would encourage you to read that post before continuing on.
The hardest thing about pregnancy has been embracing my changing body. Nine months ago, I had complete control of my appearance. I could work out whenever and however I wanted. I could eat what and how I wanted. After years of being in a roller coaster relationship, full of ups and downs and loop-de-loops, with my body I would have characterized our relationship as healthy. But I've come to realize that healthy has been a euphemism for control. I like being in control. I feel good about myself when I'm in control.
Pregnancy has taken the control right out of my hands and exposed the complicated network of issues I still struggle with. As James grew, I grew, and so did the insecurities that hovered just below the surface of my controlled exterior. I'm writing without having been fixed, without having come out on the other side of these insecurities yet. I'm still in the mess, but am not without hope.
Numbers… I'm still a slave to the numbers. My pant size can make me do the happy dance and it can send me spiraling into the blues. The same goes for my weight. I dread the beginning of each OB appointment because of the numbers that will stare back at me from that electronic scale. I make a point of kicking off my shoes, shedding any extra clothing, and emptying my bladder before being weighed, just to shave off a few ounces. I know weight gain is inevitable, but it's been difficult to see numbers rise higher and higher every month.
Identity… Being pregnant is such an in-between stage. I don't have a baby in my arms yet, but I'm not my normal unhindered self. My body has been a continuing reflection of a new stage, a new role that I wasn't quite prepared for in the first place. As my waist line expanded, I grieved the loss of being young and unencumbered. I didn't realize how much I associated my body image with my season of life. My physical shape is a barrier to the image I so enjoyed embodying (adventurous, newlywed, with no strings attached). The changes in my body are ever hinting at a major life change that still scares me poopless sometimes.
Appearance… I just don't look the same. I wasn't always happy with how I looked pre pregnancy, but I definitely liked my appearance more then than now. My arms are softer and my legs have lost their tone. My love handles have grown and my face is more full. I feel unattractive and I miss my pre pregnancy physique. I've lost my edge in the beauty battle because I'm round and pregnant (utter shame here for making beauty a competition).
Future… I worry about never fitting into my pre pregnancy clothes again. What if I have to get rid of all my jeans and start from scratch because I failed at slimming back down. What if Tim doesn't find me as attractive post pregnancy as he did when we first got married? What if I don't have the will power to eat healthy and get back in shape?
I've been struggling with letting go of my non-mom identity and the physical ideal I can't hold up as a pregnant woman. When my value is in physical appearance and numbers, I will always be striving, striving to fit into the world's ideal. When I'm in that mode, I become focused on how to keep climbing the physical ladder. I obsess about being the prettiest person Tim sees every day (my perception, not his), which immediately sets me up for failure, disappointment, and a whole slew of negative self talk. All of which serves to sadden my Creator and alienate me from my husband and those I'm judging and comparing myself to.
There are so many aspects of this season that require grace, rest, contentment, and perseverance. Pregnancy is so much more than carrying around your developing child for 40 weeks. The trials go deeper than morning sickness, aches and pains. It's a mental, emotional, and spiritual trial that can only be meant to make us more like Jesus.
Thanks for letting me be candid about these struggles. I am very much in process and I realize postpartum is going to be a whole new process in itself.
What's been the hardest thing for you about pregnancy?
For me, one of the hardest things was being SO TIRED. I am normally a pretty energetic person, and it just really took me by surprise. But keeping things on a positive note, when I am pregnant my hair looks AMAZING. Normally its really thin but it gets almost kinda thick when I am pregnant and I love that. Also, strangely, my leg hair didn't grow much during my first pregnancy, which was awesome. No such luck the second time around. What's something you've enjoyed about pregnancy?
I was also surprised at the physical exhaustion factor – definitely knocks you on your bum!
I almost mentioned the hair thing in this post! My hair, and nails, are strong and healthy and grow so fast. The nail thing is especially nice for me because mine are generally peely and lame. I've also really enjoyed feeling James move around. I was at BSF the first time I recognized his movements and have loved knowing he's alive and well in his habitat when he moves.
I have a challenge for you. For the rest of your OB appointments stand on the scale backwards. The scale reads the same no matter which way you face and the doctor will tell you if there is anything wrong with your weight gain anyway so you don't need to look for yourself. See if that helps free your mind a bit.
I know it's hard now that your body is not your own, but God created our bodies the way he did to carry out this very special task, something that only we can do. Now, you will never again find me in a two piece swimsuit, but looking at the stretch marks that I loathed so much the first time around, I am reminded of the special connection that I got to have with my children before anyone else even got to meet them. Thank you for sharing your struggles with us. You are not alone!
Our bodies truly are amazing. Dwelling on that is much more healthy than my default.
Love the honesty. Here is a blog post from a friend of mine that helped ground me and give me hope when I was feeling much like you.
I stumbled upon that blog post, via Facebook I think, a month or so ago and thought it was really encouraging. Thanks for sharing it!
I may have shared this before, but I was super duper sick during my pregnancy with hypermesis gravidum(had to cut hours at work, hospital visits for fluids, etc.) I am working on a post about it because it was a difficult experience, but I don't want to sound whiny when I got a beautiful baby boy out of the deal. That was definitely the hardest physical part. The hardest emotional part was probably all the worry (and still is).
Can totally relate to the body issues (especially taking your shoes off at the doctor's office ;)) ! Once I started gaining weight, and especially at the end, it would make me panic a little bit to go up to a weight I had never been before.
I recently saw a pic you posted on Instagram of your prego self and had a moment of jealousy at how thin and good you looked! At that stage I was bigger and i thought gosh she'll have no trouble post baby! I know others complements don't change how we feel about ourselves but I had to share.
Being a mom starts 40 weeks before you hold that baby and I totally get the struggle of knowing how to shift roles and wondering how things will be post baby. I appreciate you being so transparent! I'll be praying for you as you go through these next months! Keep close to Jesus and he'll help you navigate this new stage.


It will be curated to include work and activity inspired by the theme from the architectural and creative communities of Brighton and Hove and Sussex and will be open to the public daily within the Love Architecture festival. The programme will include site-specific installations, exhibitions, workshops, film, talks and events.
Threshold will be based the car park of myhotel, Brighton, and is lead by RIBA Sussex Branch with a:b:i:r architects Chalk Architecture, architectural photographer Jim Stephenson and arts consultant and curator Cara Courage.
You can follow Threshold on Twitter @ThresholdHub.
Our first night of a week's worth of events kicks off at Threshold on Wednesday with an architecture themed Pecha Kucha evening.
Pecha Kucha comes from the Japanese term for 'chit-chat'. The format was devised by two architects in Tokyo about ten years ago and features a number of speakers all presenting in one night. Each speaker is allowed 20 slides, with only 20 seconds on each slide. So, each talk only lasts 6 minutes 40 seconds. This makes for a fast paced and informal evening – anyone who has ever been to, or taken part in, one of these nights knows how much fun it is as the audience is presented with lots of interesting ideas and the speakers try to keep up with the slideshow.
As with all the Threshold events, entry is free. The talks will kickoff at 7pm, but please get there a little early. On the night, we'll be giving away the first of four special edition programmes, featuring an essay by Gem Barton.


"Almost 10 000 voters" is a documentary, presenting how does democratic process work in today's Ukraine. However the director is not focused on the works of the government or the strike on the famous Maidan, but on an average small town, where the most important decisions are made. She carefully observes locals discussing politics on the street market, kids being thought how democracy works by their teacher; and young men, dressed in uniforms, going to war, not knowing if they will return. The film is a perfect example of how small-scale story make a big impact on the audience.
Director, social activist and traveler Ulyana Osovska was born in Ukraine. She graduated in international business from Kyiv National Economic University. Ulyana Osovska was an active participant of the Ukraine's revolution of 2014 which sparked her interest in documenting the world through a camera.
She debuted as a director in 2015 with the short film "Election Agent". In the same year she has co-funded the NGO Docutoloka, which brings together artists and activists.
"Almost 10,000 Voters" (2017) is her full-length documentary debut, which held a World Premiere at the 33rd Warsaw Film Festival.


text:
Q: Pandas: reshaping dataframe by splitting columns in a column and a variable I have the following dataframe that I am trying to melt:
import numpy as np
import pandas as pd
dates = pd.date_range('1/1/2014', periods=4)
df = pd.DataFrame(np.eye(4, ), index=dates, columns=['A_var1', 'A_var2', 'B_var1', 'B_var2'])
print(df)

             A_var1  A_var2  B_var1  B_var2
2014-01-01     1.0     0.0     0.0     0.0
2014-01-02     0.0     1.0     0.0     0.0
2014-01-03     0.0     0.0     1.0     0.0
2014-01-04     0.0     0.0     0.0     1.0

I want to obtain the following:
            type    var1    var2
2014-01-01   A      1.0     0.0
2014-01-01   B      0.0     0.0
2014-01-02   A      0.0     1.0
2014-01-02   B      0.0     0.0
2014-01-03   A      0.0     0.0
2014-01-03   B      1.0     0.0
2014-01-04   A      0.0     0.0
2014-01-04   B      0.0     1.0

Any idea on how to do that efficiently? I know I can use the melt function but I can't get it to work in that context.
Many thanks,

A: You could use stack on multi-indexed columns.
In [304]: df.columns = df.columns.str.split('_', expand=True)

In [305]: df.stack(0).reset_index(1)
Out[305]:
           level_1  var1  var2
2014-01-01       A   1.0   0.0
2014-01-01       B   0.0   0.0
2014-01-02       A   0.0   1.0
2014-01-02       B   0.0   0.0
2014-01-03       A   0.0   0.0
2014-01-03       B   1.0   0.0
2014-01-04       A   0.0   0.0
2014-01-04       B   0.0   1.0

A: One option is with the pivot_longer function from pyjanitor, using the .value placeholder:
# pip install pyjanitor
import pandas as pd
import janitor

df.pivot_longer(names_to=("type", ".value"),
                names_sep="_",
                ignore_index=False,
                sort_by_appearance = True)

          type  var1  var2
2014-01-01   A   1.0   0.0
2014-01-01   B   0.0   0.0
2014-01-02   A   0.0   1.0
2014-01-02   B   0.0   0.0
2014-01-03   A   0.0   0.0
2014-01-03   B   1.0   0.0
2014-01-04   A   0.0   0.0
2014-01-04   B   0.0   1.0

The .value keeps the part of the column associated with it as header, while the rest goes into the type column.

redpajama_set_name: RedPajamaStackExchange


March 26, 2019 Belmont at Southern Ill.
Vasic, Nikola reached first on a fielding error by 3b.
Epps, Grey singled to right field, RBI; Vasic, Nikola scored, unearned.
Crider hit by pitch; Gardner advanced to second.
Hudson, Brad singled, bunt, advanced to second on a throwing error by 3b; Fugitt, Addison advanced to third.
Hudson, Brad advanced to third on a wild pitch; Fugitt, Addison scored on a wild pitch, unearned.
Holland doubled to left field.
Archer, Philip struck out swinging, reached first on a throwing error by c.
Weber, J.T. singled to right center; Archer, Philip advanced to second.
Lyon, Alex singled to right field; Weber, J.T. advanced to second; Archer, Philip advanced to third.
Butkiewicz, Colin flied out to cf, sacrifice fly, RBI; Archer, Philip scored, unearned.
Marcotte to p for Hubbell.
Liskey to p for Marcotte.
Weber, J.T. reached first on a fielding error by 2b, advanced to second; Archer, Philip advanced to third.
Lyon, Alex doubled to right center, 2 RBI; Weber, J.T. scored, unearned; Archer, Philip scored.
Klotz to p for Liskey.
Hudson, Brad tripled, scored on a throwing error by ss, 2 RBI, unearned; Fugitt, Addison scored, unearned; Lyon, Alex scored, unearned.
Boeckmann, Henry to p for Steidl, Matthew.
Crider walked; Gardner advanced to second.
Ehrhart to 2b for Shacklett.
DeSomer, Matt to p for Boeckmann, Henry.
Ehrhart singled to right field; Capobianco advanced to third.
Taylor reached first on a fielding error by 2b; Ehrhart advanced to second; Capobianco scored, unearned.
Fowlkes to p for Klotz.
Lyon, Alex singled up the middle.
Crawford, Kenton to rf for Vasic, Nikola.
Mitson, Corey to p for DeSomer, Matt.
Severance advanced to third on a wild pitch.
Crider doubled to left field, RBI; Severance scored.
Jarvis homered to right center, RBI; Crider scored.
Crawford, Kenton walked; Hudson, Brad advanced to third on a wild pitch.
Crawford, Kenton advanced to second on an error by 1b, advanced to third; Hudson, Brad scored.
Walters, Ian flied out to cf, sacrifice fly, RBI; Crawford, Kenton scored, unearned.
McDaniel, Trey to p for Mitson, Corey.
Hadden pinch hit for Adkins.


text:
Home » USDA to provide $4 million for honey bee habitat
USDA to provide $4 million for honey bee habitat
10/29/14 3:52 PM By Agri-Pulse staff
WASHINGTON, Oct.29, 2014 – Agriculture Secretary Tom Vilsack announced today that more than $4 million in technical and financial assistance will be provided to help farmers and ranchers in the Midwest improve the health of honey bees. USDA's Natural Resources Conservation Service (NRCS) is focusing the effort on five Midwestern states: Michigan, Minnesota, North Dakota, South Dakota and Wisconsin.
"The future of America's food supply depends on honey bees, and this effort is one way USDA is helping improve the health of honey bee populations," Vilsack said. "Significant progress has been made in understanding the factors that are associated with Colony Collapse Disorder and the overall health of honey bees, and this funding will allow us to work with farmers and ranchers to apply that knowledge over a broader area."
An estimated $15 billion worth of crops is pollinated by honey bees, including more than 130 fruits and vegetables, according to USDA. Today's announcement expands a successful $3 million pilot investment that was announced earlier this year.
Funding will be provided to producers through the Environmental Quality Incentives Program (EQIP) and applications are due Friday, November 21.
USDA announces $29 million to provide housing for farmworkers
USDA and EPA release report on honey bee health
USDA announces action during drought opened 2.8 million acres to haying and grazing, brought nearly $200 million in forage for producers
From June to September, the Midwest is home to more than 65 percent of the commercially managed honey bees in the country.
The USDA funding is intended to help farmers and ranchers implement conservation practices that will provide diverse food sources for honey bees. USDA noted that appropriate cover crops or rangeland and pasture management may provide a benefit to producers by reducing erosion, increasing the health of their soil, inhibiting invasive species, and providing quality forage and habitat for honey bees and other pollinators.
For more news, go to www.agri-pulse.com

redpajama_set_name: RedPajamaCommonCrawl


text:
Elmwood, WI (View All Cities)
Central (6:05am)
44.8, -92.2
Elmwood, WI
ZIP code 54740 is located in western Wisconsin and covers a slightly higher than average land area compared to other ZIP codes in the United States. It also has a slightly less than average population density.
The people living in ZIP code 54740 are primarily white. The number of middle aged adults is extremely large while the number of people in their late 20s to early 40s is large. There are also a small number of single parents and a slightly higher than average number of families. The percentage of children under 18 living in the 54740 ZIP code is slightly less than average compared to other areas of the country.
23 people per sq mi
Male 59 52 68 72 55 79 59 59 68 84 94 77 78 53 39 29 21 22
Female 57 62 60 57 47 60 52 71 47 73 82 65 62 56 33 35 34 35
Total 116 114 128 129 102 139 111 130 115 157 176 142 140 109 72 64 55 57
5-9 52 62 114
Owner 11 71 109 162 147 87 56 24
Renter 13 41 22 29 19 22 16 7
Total 24 112 131 191 166 109 72 31
Male 11 12 13 7 18 10 11 6 7 15 11 20 10 12 18 15 17 15 7 9
Female 4 11 10 16 16 7 18 13 8 15 14 14 8 9 15 9 12 12 9 9
ZIP code 54740 has a slightly less than average percentage of vacancies. The Census also indicates that there are one or more nursing homes nearby.
The majority of household are owned or have a mortgage. Homes in ZIP code 54740 were primarily built in 1939 or earlier. Looking at 54740 real estate data, the median home value of $150,200 is slightly less than average compared to the rest of the country. It is also slightly less than average compared to nearby ZIP codes. 54740 could be an area to look for cheap housing compared to surrounding areas. Rentals in 54740 are most commonly 3+ bedrooms. The rent for 3+ bedrooms is normally $500-$749/month including utilities. Prices for rental property include ZIP code 54740 apartments, townhouses, and homes that are primary residences.
For more information, see Elmwood, WI rent prices.
As with most parts of the country, vehicles are the most common form of transportation to places of employment. In most parts of the country, the majority of commuters get to work in under half an hour. However, that is not the case in 54740. It has a much lower than average number of people that make it to work in under half an hour. It is much more common for employees to have to travel over 45 minutes to their place of employment than most parts of country which could indicate bad traffic or that people typically live far from where they work.
Compared to other ZIPs, a much higher percentage of the population has received a college degree.
ZIP Code 54740 is in the Elmwood School District. There are 3 different elementary schools and high schools with mailing addresses in ZIP code 54740.
Elmwood Middle School
213 S Scott St
Elmwood, WI 54740
District: Elmwood School District
Elmwood High School
Grade Level: High/Secondary
Ellsworth, WI
Arkansaw, WI
Eau Galle, WI
Maiden Rock, WI
Menomonie, WI
Plum City, WI
Spring Valley, WI


Custom wall graphics murals printed wallpaper buena park ca 90621 is just one of the many collections of pictures or photos that are on this website. Custom wall graphics murals printed wallpaper buena park ca 90621 is posted on the category Wallpaper Design Companies in the dwigallery.com website. This post of "custom wall graphics murals printed wallpaper buena park ca 90621" was published on 08-08-2018 by paul and has been viewed 880,489 times.
We hope you can find what you need here. We always effort to show a picture with HD resolution or at least with perfect images. Custom wall graphics murals printed wallpaper buena park ca 90621 can be beneficial inspiration for those who seek an image according specific categories, you can find it in this site. Finally all pictures we have been displayed in this site will inspire you all. Thank you for visiting.
Night sky photo wallpaper galaxy wallpaper custom 3d clouds stars wall murals kids girls is posted in the category 3D Wallpaper For Wall and published on 02-02-2018 by dave. Image source: pinterest.com. Click on the title to see more..
3d photo wallpaper nature park tree murals bedroom living room sofa tv background wall mural is posted in the category 3D Wallpaper For Wall and published on 05-06-2018 by admin. Image source: pinterest.com. Click on the title to see more..
Custom wallpaper custom wall murals megaprint is posted in the category Wallpaper For Home Wall and published on 02-07-2018 by admin. Image source: megaprint.com. Click on the title to see more..
Custom wall murals photo wallpaper redcliffe imaging is posted in the category Wallpaper For Office Wall and published on 13-04-2018 by paul. Image source: custom-wallpaper-printing.co.uk. Click on the title to see more..
Custom wall graphics murals printed wallpaper buena park ca 90621 is posted in the category Wallpaper Design Companies and published on 08-08-2018 by dave. Image source: superiorsignsandgraphics.com. Click on the title to see more..


Mnemo is an integrated system to support human biographical memory. Mnemo is directed to serve people with impaired memory (e.g. Alzheimer's patients) by providing intuitive ways to benefit from large amounts of personal data.


Items deemed non-essential would include all ecommerce shipments, an inclusion which some saw as another swipe by Trump at Amazon and CEO Jeff Bezos, but the USPS denies any one company was targeted.
The task force is also recommending the USPS set up separate balance sheets for its package and mail divisions to prevent so-called "cross-subsidization" that competitors such as UPS and FedEx claim allows the USPS to set parcel pricing at below-market rates. The task force also supported continuing the government-supported package delivery service "at least in the short term" to benefit small and medium-sized ecommerce sellers in particular. This leaves the door open to a future spinoff into privatization, which many oppose.
While the task force recommended recalculating required payments to the retiree health benefits plan based on the population of employees at or near retirement age, it did not recommend ending the mandated prefunding of those benefits. Created by the 2006 Postal Accountability and Enhancement Act, the mandate has been blamed by both members of Congress and federal unions as the prime culprit in the USPS's current fiscal woes, contributing to billions in red ink.
The union representing postal workers took a dim view of the task force report, according to Federal Times. "This poorly conceived report makes many of its recommendations based on myth and misinformation that instead of improving mail services, would deliver higher prices and less service for the public," said Mark Dimondstein, president of the American Postal Workers Union, in a statement to the publication.


text:
21 Mar 2021 6:18 PM +00:00 UTC
Justice League: Zack Snyder Regrets Not Setting Up Ray Fisher's Cyborg Movie
By Nobelle Borines
There is little doubt that Zack Snyder's Justice League gave Victor Stone a much bigger role that...
There is little doubt that Zack Snyder's Justice League gave Victor Stone a much bigger role that would give the character a huge push for his solo film. However, the director admits that he wishes he did more in the Snyder Cut to truly set up the Cyborg movie starring Ray Fisher.
Snyder recently spoke to Esquire where he discussed the director's cut of Justice League. At one point, the filmmaker stated that he should have actively prepared fans for the arrival of a Cyborg movie.
"I'm very sad that I didn't push harder to set up the Cyborg movie before we shot Justice League because I'd done a lot of work to set up Wonder Woman and Aquaman and to get those movies on their feet," Snyder said. "And I really wish I had done that with Cyborg because he's really on his feet at the end of the movie. He's really ready to go."
But what convinced Snyder that it wasn't necessary in the first place? The Batman v Superman: Dawn of Justice director says he was so sure that the Cyborg film will be happening.
"I just thought, well, the truth is I just assumed that it was a no brainer that a Cyborg movie would be in the works. You know what I mean? It seems like so obvious," Snyder said.
Interestingly, the Snyder Cut concluded in a way that would set up a Cyborg movie as the next DC Extended Universe flick. However, that might not be Warner Bros.' plan anymore. After all, Fisher has already been fired from The Flash and it is unlikely that he will be brought back to star in his own film.
Although the Cyborg movie might not be happening anymore, the release of the Snyder Cut has led to fans crying out to see more of Victor Stone. There hasn't been a hashtag started yet but there is little doubt that people will be storming social media to demand the Cyborg film.
For now, Warner Bros. has not made an official announcement about Cyborg's solo movie. In the meantime, fans can see Vic Stone in action in Zack Snyder's Justice League, which is available on HBO Max and various streaming services around the globe.
Related: Zack Snyder's Justice League Black and White Edition Announced With New Teaser
The Flash: DC Comics Set to Release Three-Issue Prequel Series to the Upcoming Film
The Batman: Robert Pattinson Clarifies Controversial Comments Regarding His Work Out


Moisture in your home or business is a problem as soon as it occurs. It may develop suddenly, or gradually, but as soon as you become aware of excessive moisture building up in any area of the home, it is vital that you have it mitigated before it destroys the structure of the building and becomes costly to repair. You may need a moisture control appointment for crawl space moisture, basement moisture, or attic moisture. Wherever it is occurring, AdvantaClean of Ellis County and Waco is trained to determine the source and mitigate the problem. Moisture issues are especially damaging to buildings when they cause mold to grow. Mold needs moisture to grow and doesn't need sunlight to live. This makes it a prevalent problem in dark and damp areas.
WHAT DOES CRAWLSPACE MOISTURE LOOK LIKE?
A musty odor is wafting upward into the home.
Crawlspace moisture can be caused by leaking HVAC systems or ducts, rain water entering the crawlspace during a storm, or even animal urination. Crawlspace moisture is especially common if there is an exposed dirt floor. All of these moisture issues can quickly lead to damages inside your Waco home and mold growth. Resolve the problem swiftly, and save yourself the headache of crawlspace moisture.
ARE YOU CONCERNED ABOUT BASEMENT MOISTURE?
Basement moisture can be difficult to deal with, as most basements are unfinished and below ground level. The first step is always finding the cause of the basement moisture; fortunately, our technicians are skilled at diagnosing the source.
Attic moisture often goes unnoticed, as they are out of the way and mostly unthought of. Typically used as storage areas, attics are rarely given the proper cleaning and maintenance they need to stay healthy. If it's been a while since you've inspected the attic, it may be worth your while to take a peek. Attic moisture is very serious and often leads to mold growth.
Attics are more than just convenient storage spaces. Many attics also house portions of your HVAC system and the majority of your home's insulation.
If you're concerned about moisture issues in your Waco attic, call AdvantaClean (972) 776-2768 today to schedule a moisture control appointment . We will keep you informed of our findings every step of the way, and offer the best solutions to your problem, including mold remediation if needed.


Making their first significant offseason move to help the offense, the Washington Redskins drafted lineman Brandon Scherff with the No. 5 pick in the first round of the NFL draft Thursday night.
Scherff, listed at 6-foot-5 and about 320 pounds, is new general manager Scot McCloughan's first draft pick with the Redskins, who went 4-12 last season to finish last in the NFC East for the sixth time in seven years.
Scherff won the Outland Trophy in college as a tackle, but could be shifted to guard in the NFL. He is considered quite talented as a run blocker but might need to improve in pass protection.
The Redskins are expected to use him on the right side of their offensive line.
Leading up to the draft, it was thought the Redskins might trade down from the fifth overall spot in order to acquire extra picks. McCloughan said Monday that he'd ``love to get 10-plus'' choices, instead of the seven he started with.
But he held onto the fifth pick and got the sort of player who Washington hopes will wind up teaming with left tackle Trent Williams, taken No. 4 overall in 2000, to fend off opposing defenders.
This was the first time the Redskins had drafted a player in the first round since using the No. 2 overall selection in 2012 to get quarterback Robert Griffin III. That pick was obtained in a trade with the St. Louis Rams that cost the Redskins a bevy of choices, including first-rounders in 2013 and 2014.
Griffin led the Redskins to the 2012 NFC East title and was the NFL Offensive Rookie of the Year, but he tore knee ligaments in a playoff loss to Seattle that season. He's been in and out of the lineup since, because of injuries and coaches' decisions, but McCloughan announced this week that the Redskins planned to exercise their fifth-year contract option on the quarterback for 2016.
Until now, most of Washington's offseason changes came on defense.
Joe Barry was hired as the defensive coordinator, replacing Jim Haslett, and McCloughan brought in linemen Terrance ``Pot Roast'' Knighton, Stephen Paea and Ricky Jean-Francois, cornerback Chris Culliver, and safeties Jeron Johnson and Dashon Goldson.


text:
Michael J. Ware (born 1939, Bromley)  is a chemist and photographer, known for his work in alternative photographic processes, earlier methods of printing photographic images that were succeeded by the more common silver-gelatin used today. In the Present, Ware acts as a consultant, most recently on the history and development of the platinotype and palladium processes. His has also written about chemistry's influence on the history of photography.

Early life and education
Ware was born in 1939 in Bromley. He earned his Ph.D. at the University of Oxford in 1965. His thesis was The vibrational spectra of some inorganic complexes.

Awards and honours
 1982 Fellow of the Royal Society of Chemistry
 1990 Hood Medal, awarded by the Royal Photographic Society

Selected publications

Academic works

Published books

External links
 Mike Ware Alternative Photography

References

1939 births
Living people
Photographers from London
People from Bromley
Fellows of the Royal Society of Chemistry
Alumni of the University of Oxford

redpajama_set_name: RedPajamaWikipedia


Golden Calf Company is proud to announce a new colostrum thaw and pasteurizing unit: Calf Hero™ 6. With the capacity to process six meals of colostrum at once, it is designed for growing dairies.
This unit is the latest addition to the very successful Calf Hero™ colostrum management line, which offers state of the art technology. Built in Wisconsin of heavy duty stainless steel, it is designed with durability in mind. Exceptional user experience is guaranteed with a large 6" touch screen, oversized icons, instructions in multiple languages and an audio and visual alarm. The Calf Hero™ offers text messaging, emailing, and remote troubleshooting. SMART technology makes maintenance easy and ensures minimal downtime. The Calf Hero™ 6 even includes a cleaning cycle and a supplies ordering module.
The Calf Hero™ 6 is part of a colostrum management system that includes a full suite of accessories including colostrum testing, storage bags, freezer, and feeding tools.
This colostrum management system clearly defines everyone's responsibilities, is easy to implement, and provides fool-proof steps to assure everyday repeatability by multiple employees. The system thus enables the maternity workers to treat each calf as if it is the only calf that day.
In full-filling Golden Calf Company's mission "to provide dairy producers with innovative products for calf care", our goal is to supply dairy farms the right tools to achieve a consistently healthy calf population. Based in Bloomer, WI, Golden Calf Company has been an innovator in the dairy colostrum feeding field since 2010.