Abstract
A wide range of single- and multi-cellular parasites infect humans and other animals, causing some of the most prevalent and debilitating diseases on the planet. There have been virtually no published studies on the TRP channels of this diverse group of organisms. However, since many parasite genomes have been sequenced, it is simple to demonstrate that they are present in all parasitic metazoans and that sequences related to the yeast trp are present in many protozoans, including all the kinetoplastids. We compared the TRP genes of three species of animal and plant parasitic nematode to those of C. elegans and found that the parasitic species all had fewer such genes. These differences may reflect the phylogenetic distance between the species studied, or may be due to loss of specific gene functions following the evolution of the parasitic lifestyle. Other helminth groups, the trematodes and cestodes, seem to possess many TRPC and TRPM genes, but lack TRPV and TRPN. Most ectoparasites are insects or arachnids. We compared the TRP genes of a plant parasitic aphid and an animal parasite louse and tick with those of Drosophila. Again, all the parasitic species seemed to have fewer types of TRP channel, though the difference was less marked than for the nematodes. The aphid lacks TRPP and TRPML channel genes, whereas the tick lacked those encoding TRPVs. Again, these differences may reflect adaptation to parasitism, and could enable TRP channels to be targeted in the development of novel antiparasitic drugs.
Access provided by Autonomous University of Puebla. Download chapter PDF
Similar content being viewed by others
Keywords
20.1 Introduction
The term “parasite”, though fairly easy to define in biological terms, presents a major problem for chapters such as this as it covers a wide variety of single- and multi-celled organisms which cover a large phylogenetic range. This is illustrated by Table 20.1, which describes the variety of species that specifically parasitise humans and other mammals. This diversity of phylogeny is matched by an equal diversity in biology and life-cycles, all of which combine to make the task of producing an authoritative chapter on TRP channels across parasitology a major challenge. However, this task can be approached via the realization that many parasites are related to well-studied model organisms, and the dramatic increase in genome sequence information means that it is possible to compare the members of the TRP family encoded by parasites with those of their model cousins and perhaps even deduce what some of the physiological roles of the channels might be. For example, several of the parasites in Table 20.1 are nematodes, and the genome of the model nematode Caenorhabditis elegans was not only the first to be sequenced of any metazoan [1], it has also been superbly annotated.
There is no denying, though, that there have been hardly any investigations into the functions of TRP channels in parasites. This is a pity, and a lacuna in our knowledge that should be addressed. TRP channels present attractive drug targets [2], especially as many existing anti-parasitic compounds, especially anthelmintics, act on other ion channels [3–6], including ligand-gated ion channels and the SLO-1 potassium channels, and an improved pipeline of new drugs is always needed [7–9]. In addition, the involvement of the TRP channels in so many sensory processes implies that they will have roles in mediating the many host:parasite interactions that are essential for the completion of sometimes very complex life-cycles. Understanding these roles could dramatically improve our understanding of the molecular and cellular mechanisms that underpin the adaptations necessary for successful parasitism to evolve.
This hugely diverse grouping of parasites possesses an equal diverse collection of life-cycles. These life-cycles may be direct, involving only a single host, or indirect, requiring both a definitive host, in which sexual reproduction (if it occurs) takes place, and an intermediate host, which allows for the parasites to be dispersed and in which other developmental stages may take place. As an example, for malaria the human is a definitive host and the mosquito the intermediate host, and there are many different developmental stages within both. Even in those parasites with direct life cycles, many stages of development may take place outside the host and in the environment, for example hookworm eggs are shed into environment, then hatch and undergo several moults before reinfecting their mammalian host. A full description of the events within the various hosts is well beyond the scope of this chapter, as all parasites go through multiple developmental stages and have to find their specific niche within the host body. Many migrate around; the intestinal nematode Ascaris lumbricoides migrates from gut to liver to lungs and back to the gut during the various stages of its development within infected humans.
This requirement for replication within an animal host has severely limited the development of tools to study parasites. Some parasitic protozoans can be cultured in the laboratory and for these advanced molecular genetic tools such as gene knock-outs have been developed [10–13]. However, no such culture methods or tools are readily available for the parasitic helminths, and this has had a severely limiting effect on research on these organisms. For the nematodes and many ectoparasites, good model organisms (C. elegans and Drosophila melanogaster, respectively) are available for comparison and, at the genomic level, these comparisons have been made facile by the amount of sequence information that is now available. A considerable amount of excellent science has been carried out on the TRP channels of these model organisms ( [14–17]), but the biological relevance of these studies to parasitic species remains to be determined. Studies on other nematode gene families show that the composition of these families, and possibly their functions, differ greatly between C. elegans and parasitic species [18, 19] and so simple extrapolation may be misleading.
The biological diversity of parasites demands that this chapter be sub-divided, and we have chosen to do this along the lines of Table 20.1. Separate sections will discuss endoparasites, those organisms that invade their hosts and live inside them (which include the protozoan parasites and the helminthes, or “worms”) and the ectoparasites, organisms that live on the outside of the host, but are dependent on them for food – many are blood feeders.
20.2 Endoparasites
20.2.1 Protozoa
A very diverse group of single-celled organisms has evolved to parasitise humans and other animals. The apicomplexans, which include the Plasmodium species that cause malaria, arguably the most important infectious disease on the planet [20], are an exclusively parasitic grouping that possess a unique organelle, the apicoplast, a non-photosynthetic plastid [21], and an apical complex required for entry into host cells [22]. The kinetoplastids, which cause sleeping sickness and leishmaniasis, are a group of flagellates defined by the presence of a kinetoplast, a granule within the mitochondrion that contains DNA [23]. Both of these groups are normally transmitted via an invertebrate vector, such as a mosquito or biting fly. The metamonads are anaerobic flagellates, most of which are symbiotic with their hosts, but some of which are parasitic. A few species of amoeba are also parasitic.
Two TRPML-like genes, lmmlA (LmjF07.0910) and lmmlB (LmjF26.0990), have been described from Leishmania major, a kinetoplastid [24]. Though lmmlA is constitutively expressed throughout the life-cycle, lmmlB expression was reported to be up-regulated in amastigotes, the form of the parasite found in the vertebrate host [24]. A simple search of TrTrypDB (http://tritrypdb.org/tritrypdb/) revealed that both of these genes are conserved throughout the kinetoplastids, with similar sequences being present in the other Leishmania and Trypanosoma species. There are no published studies on the functions of these channels.
There are no other reports of TRP channels from any of these parasites, so we undertook a simple bioinformatics search of some of the available genome sequences, using the membrane-spanning domains of the yeast trp sequence [25] and of C. elegans CUP-5 (a TRPML [26]) as probes. This search revealed sequences in Trichomonas vaginalis (a metamonad) with low levels of identity (~22%) to both the yeast trp and C. elegans CUP-5, and a similar sequence (24% identity to CUP-5) in Cryptosporidium parva (an apicomplexan). The C. parvum protein (cdg6_1510) possesses membrane-spanning regions and is conserved in C. hominis and C. muris. The T. vaginalis “hits” were less convincing, and probably do not represent true channel proteins. Though none of these proteins, including those from the kinetoplastids, has yet been shown to form a genuine ion channel, these early data do suggest that that TRP-like channels might be present in a wide variety of protozoan parasites. We found no evidence of any sequence similar to other sub-families of TRPs, except for TRPML, in these organisms, nor did we find any evidence of TRP-like channels in Plasmodium falciparum, Toxoplasma gondii, Neospora caninum or Giardia lamblia. This may be because these organisms do not have any such genes, or it might be that they are just too divergent to be easily detected – further work is clearly needed to clarify this. Apicomplexans possess most of the other components for regulation of intracellular [Ca2+] found in other eukaryotes, though P. falciparum and C. parva apparently lack voltage-operated Ca2+ channels or PMCA [27]. Given the role of TRPML channels in intracellular events in yeast [25] and higher eukaryotes [28, 29], we would hypothesise that any such channels in protozoan parasites will be involved in the intracellular trafficking and membrane fusion events that take place in these organisms [30, 31]. One possible example of such an event is the fusion of acidocalcisomes, a dense acidic organelle, to the contractile vacuole of T. cruzi and related organisms that is part of the osmoregulation process [32]. Additionally, entry into and replication within host cells by the kinetoplastids is intimately associated with lysosomal-like organelles and membrane fusion events [33, 34]. These events are regulated by TRPML in uninfected cells [28] and it is possible that the related proteins encoded by these parasites function as part of the process by which they control this compartment in their hosts.
20.2.2 Helminths
20.2.2.1 Nematodes
Nematodes, or roundworms, represent about 80% of all animal species. Many of these species are parasitic, causing chronic infections and debilitating, though rarely fatal, disease. A couple of parasitic genome sequences have been published [35, 36] in addition to that of the “model worm”, C. elegans [1], and a considerable amount of sequence information is available, primarily from the groups at Washington University, St Louis, USA (http://www.nematode.net) [37] and the Sanger Centre, Cambridge, U.K. (http://www.sanger.ac.uk/Projects/Helminths/). Many of the C. elegans TRP genes have been extensively studied and reviewed [17]; some of this information is summarized in Table 20.2. However, phylogenetic analysis suggest that C. elegans may not be typical of all nematodes [38], and previous analyses have revealed big differences between the ion channel genes of C. elegans and some parasitic species [19]. We searched the annotated genes of the parasitic nematode Brugia malayi [35] and the plant parasite Meloidogyne incognita [36] together with the unannotated sequence of Trichinella spiralis (http://genome.wustl.edu/genomes/view/trichinella_spiralis/) for TRP channel sequences; the results are summarized in Table 20.2. Compared to C. elegans, it seems that all three parasite species possess fewer such genes, as observed previously for the ligand-gated ion channels [19]; this may not reflect any physiological simplicity of the parasites, but rather the number of gene duplications that have occurred during C. elegans evolution, though considerably more comparative genomic analysis is required before any firm conclusions can be drawn. We also searched the B. malayi genome (http://blast.jcvi.org/er-blast/index.cgi?project=bma1) for TRPM-, TRPA- and TRPP-related sequences in case these had not been annotated; we found some evidence of TRPM-like sequences but not for TRPA or TRPP. Many of the TRP channels are involved in sensory processes, in nematodes as in other organisms, and the loss of these channels may reflect a reduction in the complexity of sensory inputs encountered by the animal parasites as they inhabit the homeostatically maintained environments of their vertebrate and, in the case of B. malayi, invertebrate hosts. The T. spiralis and M. incognita genomes contain members of most of the TRP channel sub-families, but not TRPA, nor TRPP; M. incognita seems to have more TRPM channels than the animal parasites. The TRPP channels of C. elegans are required in males for successful mating and responses to hermaphrodite pheromones [39, 40]; their absence from the parasitic species may reflect differences in mating behavior between the species. This hypothesis may be supported by the absence of an obvious spe-41 from B. malayi and the divergent form of this gene seen in T. spiralis since this gene is also involved in mating [41]. The TRPV channels of C. elegans function as heteromers [42], and the reduction in TRPV gene number in the parasites may reflect an expression of homomeric channels in these species.
20.2.2.2 Trematodes and Cestodes
The most important trematode, or flatworm, parasites of humans are the schistosomes, or blood flukes [20]. There are three major species of these, S. mansoni, S. japomonicum and S. haematobium: we searched the completed and annotated genome of S. mansoni [43] for TRP channel genes (Table 20.3). We also searched the incomplete and unannotated genome of the tapeworm (cestode) Echinococcus multilocularis (http://www.sanger.ac.uk/Projects/Echinococcus/) for TRP channel sequences. These searches produced similar and quite remarkable results; both organisms possess multiple TRPC and TRPM genes (at least nine in S. mansoni), single TRPA, TRPP and TRPML genes, and apparently no TRPV or TRPN genes. To date, there is no information of the function of any of these channels; there are ESTs for several of the S. mansoni TRP genes, but these seem to be expressed at low levels; Sm_169150 (a TRPC) and Sm_147140 (a TRPM) are the most highly represented, with Sm_147140 having multiple ESTs from adult libraries. In mammals and nematodes, TRPM6 and TRPM7 channels are required for ion homeostasis in the gut [44–46]. Since trematodes and cestodes absorb most of their nutrients, including ions, from the blood or gut contents of the host organisms, it is tempting to speculate that one role of the parasite TRPMs might be in the uptake or release of essential ions. The apparent absence of TRPV channels from these organisms might, as with the parasitic nematodes, reflect the simpler sensory requirements of a parasitic life-style.
20.3 Ectoparasites
“Ectoparasites” is a term that makes biological but very little phylogenetic sense. For the purposes of this chapter, we have chosen to focus on a few examples of insect and arachnid parasites, the bugs, lice and ticks. Bugs, such as bedbugs, are closely related to aphids, which could be considered to be ectoparasites of plants; the genome sequence of the pea aphid, Acyrthosiphon pisum, has just been published [47], resulting in an analysis of the ion channel genes, including the TRP channels, present in this organism [48]. We compared the predicted TRP channel genes in A. pisum and D. melanogaster to those annotated from the human head louse, Pediculus humanus, and the tick, Ixodes scapularis (Table 20.4). We also carried out some Blast searches using the Drosophila peptide sequences as probes. The results, which may be incomplete, show a remarkable conservation in gene number between the various insects and ticks, with members of all seven sub-families of TRP channel present except for TRPV in the tick, and TRPP and TRPML in the aphid. This greater conservation of gene number between parasitic and non-parasitic insects, as compared to nematodes, may reflect the more limited adaptations required for ectoparasitism as opposed to endoparasitism. Ticks, which are not insects but arachnids, find their hosts by detecting their temperature and carbon dioxide “footprints” and so might be expected to conserve some of the temperature detecting channels of other arthropods. Insects detect heat via several TRP channels, including the TRPV channels [49], trpA1 [50] and pyrexia [51]; of these we found a clear homologue for only trpA1 (ISCW011428) in I. scapularis. TrpA1 has a role in larval thermotaxis in Drosophila [50] and so might make a good candidate for mediating the same phenomenon in ticks. Head lice move between hosts as adults and seem to have retained a larger number of temperature-sensitive TRP channels to allow them to find a new host at this stage of the life-cycle. The larvae, or nymphs, are not considered to be as infectious, which might explain the apparent loss of the larval thermotaxis gene, trpA1, in the louse.
20.4 Conclusions
An immediate conclusion from this brief survey is that, for invertebrates, far more is known about the TRP channels of model than target organisms. Even from the simple bioinformatics searches that we have carried out, it is clear that the TRP channel genes can vary, especially in the protozoa and helminths, and that these differences between species do warrant further investigation. It may be premature to consider TRPs to be viable drug targets, but they are clearly worth considering and exploring further. We hope to read about such studies in the years to come.
References
C. elegans Sequencing Consortium (1998) Genome sequence of the nematode C. elegans: a platform for investigating biology. Science 282:2012–2018
Kumari S, Kumar A, Samant M, Singh N, Dube A (2008) Discovery of Novel Vaccine Candidates and Drug Targets Against Visceral Leishmaniasis Using Proteomics and Transcriptomics. Curr Drug Targets 9:938–947
Martin RJ, Murray I, Robertson AP, Bjorn H, Sangster N (1998) Anthelmintics and ion-channels: after a puncture, use a patch. Int J Parasitol 28:849–862
Wolstenholme AJ, Rogers AT (2005) Glutamate-gated chloride channels and the mode of action of the avermectin/milbemycin anthelmintics. Parasitology 131:S85–S95
Harder A, Holden-Dye L, Walker RJ, Wunderlich F (2005) Mechanisms of action of emodepside. Parasitol Res 97:S1–S10
Kaminsky R, Ducray P, Jung M, Clover R, Rufener L, Bouvier J, Schorderet Weber S, Wenger A, Wieland-Berghausen S, Goebel T, Gauvry N, Pautrat F, Skripsky T, Froelich O, Komoin-Oka C, Westlund B, Sluder A, Maser P (2008) A new class of anthelmintics effective against drug-resistant nematodes. Nature 452:176–180
Escalante AA, Smith DL, Kim Y (2009) The dynamics of mutations associated with anti-malarial drug resistance in Plasmodium falciparum. Trends Parasitol 25:557–563
Geary TG, Woo K, McCarthy JS, Mackenzie CD, Horton J, Prichard RK, de Silva NR, Olliaro PL, Lazdins-Helds JK, Engels DA, Bundy DA (2010) Unresolved issues in anthelmintic pharmacology for helminthiases of humans. Int J Parasitol 40:1–13
Wilkinson SR, Kelly JM (2009) Trypanocidal drugs: mechanisms, resistance and new targets. Expert Revs Mol Med 11:e31
Clayton CE (1999) Genetic manipulation of kinetoplastida. Parasitology Today 15:372–378
Thathy V, Menard R (2002) Gene targeting in Plasmodium berghei. Methods Mol Med 72:317–331
Trager W, Jensen JB (1976) Human malaria parasites in continuous culture. Science 193: 673–675
Wu Y, Sifri CD, Lei HH, Wellems TE (1995) Transfection of Plasmodium falciparum within human red blood cells. Proc Natl Acad Sci USA 92:973–977
Benton R (2008) Chemical sensing in Drosophila. Curr Opin Neurobiol 18:357–363
Kahn-Kirby AH, Bargmann CI (2006) TRP channels in C. elegans. Ann Rev Physiol 68: 719–736
Montell C (1999) Visual transduction in Drosophila. Ann Rev Cell Dev Biol 15:231–268
Xiao R, Xu XZ (2009) Function and regulation of TRP family channels in C. elegans. Pflugers Arch 458:851–860
Ardelli BF, Stitt LE, Tompkins JB (2010) Inventory and analysis of ATP-binding cassette (ABC) systems in Brugia malayi. Parasitology 137:1195–1212
Williamson SM, Walsh TK, Wolstenholme AJ (2007) The cys-loop ligand-gated ion channel gene family of Brugia malayi and Trichinella spiralis: a comparison with Caenorhabditis elegans. Inv Neurosci 7:219–226
Mathers CD, Ezzati M, Lopez AD (2007) Measuring the burden of neglected tropical diseases: the global burden of disease framework. PLoS Negl Trop Dis 1:e114
Obornik M, Janouskovec J, Chrudimsky T, Liukes J (2009) Evolution of the apicoplast and its hosts: from heterotrophy to autotrophy and back again. Int J Parasitol 39:1–12
Blackman MJ, Bannister LH (2001) Apical organelles of Apicomplexa: biology and isolation by subcellular fractionation. Mol Biochem Parasitol 117:11–25
de Souza W, Attias M, Rodrigues JC (2009) Particularities of mitochondrial structure in parasitic protists (Apicomplexa and Kinetoplastida). Int J Biochem Cell Biol 41:2069–2080
Chenik M, Douagi F, Ben Achour Y, Ben Khalef N, Ouakad M, Louzir H, Dellagi K (2005) Characterization of two different mucolipin-like genes from Leishmania major. Parasitol Res 98:5–13
Palmer CP, Zhou XL, Lin J, Loukin SH, Kung C, Saimi Y:A (2001) TRP homolog in Saccharomyces cerevisiae forms an intracellular Ca(2+)-permeable channel in the yeast vacuolar membrane. Proc Natl Acad Sci USA 98:7801–7805
Fares H, Greenwald I (2001) Regulation of endocytosis by CUP-5, the Caenorhabditis elegans mucolipin-1 homolog. Nat Genet 28:64–68
Nagamune K, Moreno SN, Chini EN, Sibley LD (2008) Calcium regulation and signaling in apicomplexan parasites. Subcell Biochem 47:70–81
Pryor PR, Reimann F, Gribble FM, Luzio JP (2006) Mucolipin-1 is a lysosomal membrane protein required for intracellular lactosylceramide traffic. Traffic 7:1388–1398
Martina JA, Lelouvier B, Puertollano R (2009) The calcium channel mucolipin-3 is a novel regulator of trafficking along the endosomal pathway. Traffic 10:1143–1156
Cesbron-Delauw MF, Gendrin C, Travier L, Ruffiot P, Mercier C (2008) Apicomplexa in mammalian cells: trafficking to the parasitophorous vacuole. Traffic 9:657–664
Ravindran S, Boothroyd JC (2008) Secretion of proteins into host cells by Apicomplexan parasites. Traffic 9:647–656
Rohloff P, Docampo R (2008) A contractile vacuole complex is involved in osmoregulation in Trypanosoma cruzi. Exp Parasitol 118:17–24
Andrade LF, Andrews NW (2004) Lysosomal fusion is essential for the retention of Trypanosoma cruzi inside host cells. J Exp Med 200:1135–1143
Wilson J, Huynh C, Kennedy KA, Ward DM, Kaplan J, Aderem A, Andrews NW (2008) Control of parasitophorous vacuole expansion by LYST/Beige restricts the intracellular growth of Leishmania amazonensis. PLoS Pathogens 4:e100179
Ghedin E, Wang S, Spiro D, Caler E, Zhao Q, Crabtree J, Allen JE, Delcher AL, Guiliano DB, Miranda-Saavedra D, Angiuoli SV, Creasy T, Amedeo P, Haas B, El-Sayed NM, Wortman JR, Feldblyum T, Tallon L, Schatz M, Shumway M, Koo H, Salzberg SL, Schobel S, Pertea M, Pop M, White O, Barton GJ, Carlow CKS, Crawford MJ, Daub J, Dimmic MW, Estes CF, Foster JM, Ganatra M, Gregory WF, Johnson NM, Jin J, Komuniecki R, Korf I, Kumar S, Laney S, Li B-W, Li W, Lindblom TH, Lustigman S, Ma D, Maina CV, Martin DMA, McCarter JP, McReynolds L, Mitreva M, Nutman TB, Parkinson J, Peregrin-Alvarez JM, Poole C, Ren Q, Saunders L, Sluder AE, Smith K, Stanke M, Unnasch TR, Ware J, Wei AD, Weil G, Williams DJ, Zhang Y, Williams SA, Fraser-Liggett C, Slatko B, Blaxter ML, Scott AL (2007) Draft genome of the filarial nematode parasite Brugia malayi. Science 317:1756–1760
Abad P, Gouzy J, Aury JM, Castagnone-Sereno P, Danchin EGJ, Deleury E, Perfus-Barbeoch L, Anthouard V, Artiguenave F, Blok VC, Caillaud MC, Coutinho PM, Dasilva C, De Luca F, Deau F, Esquibet M, Flutre T, Goldstone JV, Hamamouch N, Hewezi T, Jaillon O, Jubin C, Leonetti P, Magliano M, Maier TR, Markov GV, McVeigh P, Pesole G, Poulain J, Robinson-Rechavi M, Sallet E, Segurens B, Steinbach D, Tytgat T, Ugarte E, van Ghelder C, Veronico P, Baum TJ, Blaxter M, Bleve-Zacheo T, Davis EL, Ewbank JJ, Favery B, Grenier E, Henrissat B, Jones JT, Laudet V, Maule AG, Quesneville H, Rosso MN, Schiex T, Smant G, Weissenbach J, Wincker P (2008) Genome sequence of the metazoan plant-parasitic nematode Meloidogyne incognita. Nat Biotechnol 26:909–915
Wylie T, Martin JC, Dante M, Mitreva MD, Clifton SW, Chinwalla A, Waterston RH, Wilson RK, McCarter JP (2004) Nematode.net: a tool for navigating sequences from parasitic and free-living nematodes. Nucleic Acids Res 32:D423–D426
Blaxter ML, De Ley P, Garey JR, Liu LX, Scheldeman P, Vierstrate A, Vanfleteren JR, Mackey LY, Dorris M, Frisse LM, Vida JT, Thomas WK (1998) A molecular evolution framework for the phylum Nematoda. Nature 392:71–75
Barr MM, DeModena J, Braun D, Nguyen CQ, Hall DH, Sternberg PW (2001) The Caenorhabditis elegans autosomal dominant polycystic kidney disease gene homologs lov-1 and pkd-2 act in the same pathway. Curr Biol 11:1341–1346
Barr MM, Sternberg PW (1999) A polycystic kidney-disease gene homologue required for male mating behaviour in C. elegans. Nature 401:386–389
Xu XZS, Sternberg PW (2003) A C. elegans sperm TRP protein required for sperm-egg interactions during fertilization. Cell 114:285–297
Jose AM, Bany IA, Chase DL, Koelle MR (2006) A specific subset of TRPV channels in Caenorhabditis elegans endocrine cells function as mixed heteromers to promote neurotransmitter release. Genetics 175:93–105
Berriman M, Haas BJ, LoVerde PT, Wilson RA, Dillon GP, Cerqueira GC, Mashiyama ST, Al-Lazikani B, Andrade LF, Ashton PD, Aslett MA, Bartholomeu DC, Blandin G, Caffrey CR, Coghlan A, Coulson R, Day TA, Delcher A, DeMarco R, Djikeng A, Eyre T, Gamble JA, Ghedin E, Gu Y, Hertz-Fowler C, Hirai H, Hirai Y, Houston R, Ivens A, Johnston DA, Lacerda D, Macedo CD, McVeigh P, Ning ZM, Oliveira G, Overington JP, Parkhill J, Pertea M, Pierce RJ, Protasio AV, Quail MA, Rajandream MA, Rogers J, Sajid M, Salzberg SL, Stanke M, Tivey AR, White O, Williams DL, Wortman J, Wu WJ, Zamanian M, Zerlotini A, Fraser-Liggett CM, Barrell BG, El-Sayed NM (2009) The genome of the blood fluke Schistosoma mansoni. Nature 460:352–365
Teramoto T, Lambie EJ, Iwasaki K (2005) Differential regulation of TRPM channels governs electrolyte homeostasis in the C. elegans intestine. Cell Metab 1:343–354
Teramoto T, Sternick LA, Kage-Nakadai E, Sajjadi S, Siembida J, Mitani S, Iwasaki K, Lambie EJ (2010) Magnesium excretion in C. elegans requires the activity of the GTL-2 TRPM channel. PLoS One 5:e9589
Schmitz C, Perraud AL, Johnson CO, Inabe K, Smith MK, Penner R, Kurosaki T, Fleig A, Scharenberg AM (2003) Regulation of vertebrate cellular Mg2+ homeostasis by TRPM7. Cell 114:191–200
The International Aphid Genomics Consortium (2010) Genome Sequence of the Pea Aphid Acyrthosiphon pisum. PLoS Biol 8:e1000313
Dale RP, Jones AK, Tamborindeguy C, Davies TGE, Amey JS, Williamson S, Wolstenholme A, Field LM, Williamson MS, Walsh TK, Sattelle DB (2010) Identification of ion channel genes in the Acyrthosiphon pisum genome. Insect Mol Biol 19(Suppl 2):141–153
Chentsova NA, Gruntenko NE, Bogomolova EV, Adonyeva NV, Karpova EK, Rauschenbach IY (2002) Stress response in Drosophila melanogaster strain inactive with decreased tyramine and octopamine contents. J Comp Physiol B 172:643–650
Rosenzweig M, Brennan KM, Taylor TD, Phelps PO, Patapoutian A, Garrity PA (2005) The Drosophila ortholog of vertebrate TRPA1 regulates thermotaxis. Genes Dev 19:419–424
Lee Y, Lee Y, Lee J, Bang S, Hyun S, Kang J, Hong ST, Bae E, Kaang BK, Kim J (2005) Pyrexia is a new thermal transient receptor potential channel endowing tolerance to high temperatures in Drosophila melanogaster. Nat Genet 37:305–310
Colbert HA, Smith TL, Bargmann CI (1997) OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. J Neurosci 17:8259–8269
Feng ZY, Li W, Ward A, Piggott BJ, Larkspur ER, Sternberg PW, Xu XZS (2006) A C. elegans model of nicotine-dependent behavior: regulation by TRP-family channels. Cell 127:621–633
White JQ, Nicholas TJ, Gritton J, Truong L, Davidson ER, Jorgensen EM (2007) The sensory circuitry for sexual attraction in C. elegans males. Curr Biol 17:1847–1857
Tobin D, Madsen D, Kahn-Kirby AH, Peckol E, Moulder G, Barstead R, Maricq A, Bargmann C (2002) Combinatorial expression of TRPV channel proteins defines their sensory functions and subcellular localization in C. elegans neurons. Neuron 35:307–318
West RJ, Sun AY, Church DL, Lambie EJ (2001) The C. elegans gon-2 gene encodes a putative TRP cation channel protein required for mitotic cell progression. Gene 266:103–110
Kwan CS, Vazquez-Manrique RP, Ly S, Goyal K, Baylis HA (2008) TRPM channels are required for rhythmicity in the ultradian defecation rhythm of C. elegans. BMC Physiol 8:11
Xing J, Yan XH, Estevez A, Strange K (2008) Highly Ca2+-selective TRPM channels regulate IP3-dependent oscillatory Ca2+ signaling in the C. elegans intestine. J Gen Physiol 131: 245–255
Kindt KS, Viswanath V, Macpherson L, Quast K, Hu H, Patapoutian A, Schafer WR (2007) Caenorhabditis elegans TRPA-1 functions in mechanosensation. Nat Neurosci 10:568–577
Kindt KS, Quast KB, Giles AC, De S, Hendrey D, Nicastro I, Rankin CH, Schafer WR (2007) Dopamine mediates context-dependent modulation of sensory plasticity in C. elegans. Neuron 55:662–676
Kim J, Chung YD, Park DY, Choi S, Shin DW, Soh H, Lee HW, Son W, Yim J, Park CS, Kernan MJ, Kim C (2003) A TRPV family ion channel required for hearing in Drosophila. Nature 424:81–84
Hamada FN, Rosenzweig M, Kang K, Pulver SR, Ghezzi A, Jegla TJ, Garrity PA (2008) An internal thermal sensor controlling temperature preference in Drosophila. Nature 454: 217–220
Tracey WD, Wilson RI, Laurent G, Benzer S (2003) painless, a Drosophila gene essential for nociception. Cell 113:261–273
Gopfert MC, Robert D (2003) Motion generation by Drosophila mechanosensory neurons. Proc Natl Acad Sci USA 100:5514–5519
Gao Z, Ruden DM, Lu X (2003) PKD2 cation channel is required for directional sperm movement and male fertility. Curr Biol 13:2175–2178
Venkatachalam K, Long AA, Elsaesser R, Nikolaeva D, Broadie K, Montell C (2008) Motor deficit in a Drosophila model of mucolipidosis type IV due to defective clearance of apoptotic cells. Cell 135:838–851
Acknowledgments
We would like to thank all of those who make their sequence information publically available. The T. spiralis sequence data were produced by the Genome Sequencing Center at Washington University School of Medicine in St Louis and can be obtained from http://genome.wustl.edu. Funding for the sequence characterization of the Trichinella genome is being provided by the National Human Genome Research Institute (NHGRI), National Institutes of Health (NIH). The E. multilocularis data were produced by the Sanger Centre, Hinxton UK and funded by the Wellcome Tust. The P. humanus and I. scapularis data were obtained from Vectorbase (http://www.vectorbase.org), an NIAID Bioinformatics Resource Center for Invertebrate Vectors of Human Pathogens.
Author information
Authors and Affiliations
Corresponding author
Editor information
Editors and Affiliations
Rights and permissions
Copyright information
© 2011 Springer Science+Business Media B.V.
About this chapter
Cite this chapter
Wolstenholme, A.J., Williamson, S.M., Reaves, B.J. (2011). TRP Channels in Parasites. In: Islam, M. (eds) Transient Receptor Potential Channels. Advances in Experimental Medicine and Biology, vol 704. Springer, Dordrecht. https://doi.org/10.1007/978-94-007-0265-3_20
Download citation
DOI: https://doi.org/10.1007/978-94-007-0265-3_20
Published:
Publisher Name: Springer, Dordrecht
Print ISBN: 978-94-007-0264-6
Online ISBN: 978-94-007-0265-3
eBook Packages: Biomedical and Life SciencesBiomedical and Life Sciences (R0)