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The Limits on Trypanosomatid Morphological Diversity

The Limits on Trypanosomatid Morphological Diversity

  • Richard John Wheeler, 
  • Eva Gluenz, 
  • Keith Gull
PLOS
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Abstract

Cell shape is one, often overlooked, way in which protozoan parasites have adapted to a variety of host and vector environments and directional transmissions between these environments. Consequently, different parasite life cycle stages have characteristic morphologies. Trypanosomatid parasites are an excellent example of this in which large morphological variations between species and life cycle stage occur, despite sharing well-conserved cytoskeletal and membranous structures. Here, using previously published reports in the literature of the morphology of 248 isolates of trypanosomatid species from different hosts, we perform a meta-analysis of the occurrence and limits on morphological diversity of different classes of trypanosomatid morphology (trypomastigote, promastigote, etc.) in the vertebrate bloodstream and invertebrate gut environments. We identified several limits on cell body length, cell body width and flagellum length diversity which can be interpreted as biomechanical limits on the capacity of the cell to attain particular dimensions. These limits differed for morphologies with and without a laterally attached flagellum which we suggest represent two morphological superclasses, the ‘juxtaform’ and ‘liberform’ superclasses. Further limits were identified consistent with a selective pressure from the mechanical properties of the vertebrate bloodstream environment; trypanosomatid size showed limits relative to host erythrocyte dimensions. This is the first comprehensive analysis of the limits of morphological diversity in any protozoan parasite, revealing the morphogenetic constraints and extrinsic selection pressures associated with the full diversity of trypanosomatid morphology.

Introduction

Protozoan parasite life cycles are often characterised by specialised proliferative and transmissive life cycle stages, each of which represents an adaptation to that host environment (for a replicative stage) or a pre-adaptation to the next host environment and any conditions likely to be encountered during transmission (a transmissive stage). It is often the case that transmissive stages are non-proliferative, meaning a parasite life cycle is often made up of several linked proliferative cycles. Trypanosomatids, which are a diverse order of exclusively parasitic protozoa with a monoxonous life cycle in an insect host or a dixenous life cycle between an invertebrate and vertebrate or plant host, include many excellent examples of this life cycle structure. This family includes the human pathogens Leishmania spp., Trypanosoma brucei and Trypanosoma cruzi.

Life cycle stage adaptation may incorporate many metabolic, biochemical and cell biological adaptations, including adaptation of cell shape. In trypanosomatids large morphological variation occurs both between life cycle stages and between species, despite great ultrastructural similarity which is universally conserved [1], [2]. This diversity of shape has been catalogued extensively as light microscopy provided the earliest means for classifying these microorganisms, with six major morphological classes commonly defined by the position and depth of the flagellar pocket, flagellum length, and lateral attachment of the flagellum to the cell body (Figure 1) [3]. The function of these cell shapes is largely unknown, although there are examples indicating that correct morphogenesis is vital for pathogenicity [4]. Together these properties mean the trypanosomatids are an excellent model for considering the function of cell shape in a unicellular parasite’s pathogenicity, and how this links to the capacity for morphological change for adaptation to different host environments.

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Figure 1. The major morphological classes of trypanosomatids.

Diagrams of six common, easily distinguished trypanosomatid morphologies [3]. A. Morphologies with a flagellum laterally attached to the cell body. B. Morphologies with a free flagellum (no lateral attachment of the flagellum to the cell body extending beyond the flagellar pocket neck). Metrics used to record cell morphology are indicated (cell body length, free flagellum length, kinetoplast–posterior distance (KP) and nucleus–posterior distance (NP)). Genera in which each morphology occurs [28] in are indicated, monophyletic genera [25] are underlined. C. Amastigote morphology, which does not have a long, motile, flagellum. D. Key. Structures associated with the flagellum (the basal body/pro-basal body pair (BB), flagellar pocket (FP), Axoneme (Ax) and paraflagellar rod (PFR)) are indicated.

https://doi.org/10.1371/journal.pone.0079581.g001

The diversity, and limits on diversity, of the morphologies of life cycle stages of any parasite are likely to have arisen through two biological phenomena. Firstly intrinsic biomechanical constraints arising from the cell organisation (and its growth and division) may limit the range of potential viable cell shapes. Secondly selective pressures from the host environment may limit the range of these viable cell shapes actually observed in different host environment. In qualitative terms if occurrence of particular morphological classes are limited to particular environments it is suggestive of a selection pressure; for example in trypanosomatids a selection for trypomastigotes in the vertebrate bloodstream [1], [5][9] and for amastigotes in intracellular life cycle stages [10][12] appear to have occurred. In a similar line of reasoning, trypanosomatid morphologies universally have a sub-pellicular microtubule cytoskeleton, flagellum and flagellar pocket, even in immotile amastigotes [10][14], indicating their role in basic cell organisation confers a biomechanical constraint. This limit in basic cell organisation is consistent with the well known and diverse roles of the flagellar pocket and associated structures in trypanosomatids, particularly for kinetoplast division [15], [16] and endo and exocytosis [17][20] amongst others [21].

We reasoned that more information about constraints and selective pressures acting on trypanosomatid shape could be inferred by analysing limits on trypanosomatid shape and size quantitatively, and that this could be done most effectively if we considered multiple classes of trypanosomatid morphology in combination. Whether any particular limit in morphological diversity arises from a constraint or a selective pressure could be determined by analysing multiple classes of morphology in different environments. Constraints universal to all trypanosomatids will give rise to limits to cell shape in all species and classes, while constraints restricted to only particular morphological classes will give rise to limits to cell shape within only those classes. Limits in cell shape observed in trypanosomatids of a particular morphological class in one host environment but not another may have arisen through host selective pressures.

In order to detect these potential constraints or selective pressures, we performed a literature-based meta-analysis of trypanosomatid morphology in a wide range of species. Morphometric data from 248 isolates of trypanosomatids from the vertebrate bloodstream or invertebrate host were extracted from the literature to analyse cell length, cell width, flagellum length and flagellar pocket positioning. We established the concept of two morphological superclasses, the juxtaforms (trypomastigotes and epimastigotes) and liberforms (promastigotes, choanomastigotes and opisthomastigotes) on the bases of the phylogeny of the species in which they occur and the incapacity of trypanosomatids to transition between these two superclasses in the life cycle as a basis for analysing these morphometric data. The correlations of quantitative morphological measures were used to analyse whether morphological superclasses are a valid concept and determine limits on morphological variation which may be associated with constraints intrinsic to the juxtaforms and liberforms. In juxtaforms, morphological diversity, in combination with correlation of trypanosomatid shape with host erythrocyte sizes, was used to analyse limits on morphological variation in the blood environment. Some of these limits may be associated with selective pressures from the bloodstream and we determine which are consistent with proposed functions of cell shape and motility in T. brucei. We then discuss which mechanisms may apply constraints and selection pressures consistent with our observations which may have driven the evolution of trypanosomatid morphological diversity.

Results

In order to perform the meta-analysis of limits to trypanosomatid cell shape associated with constraints or selective pressures we generated a database of previously published trypanosomatid morphometric data for motile life cycle stages of many different species from different vertebrate and invertebrate hosts (Table S1). The literature describing new species or isolates was systematically surveyed, guided by existing indices of trypanosomes [22], [23] and other trypanosomatids [23], [24], and supplemented by database searches for more recently described species and isolates. This yielded approximately 250 references and each was screened to determine if it included suitable morphometric data. Briefly (for more detail see Materials and Methods), if morphology measurements were available and taken from a sample immediately derived from a host or axenic cultures immediately derived from an isolate from a host, and data were presented as representative of the complete range of trypanosomatid dimensions present in that life cycle stage then it was judged to be of sufficient quality for inclusion in the morphometry database. Cell body length and width, free and total flagellum length and kinetoplast to posterior distance and the ranges of each of these measures (or the available subset) were recorded. All morphological analyses, unless otherwise indicated, are derived from these data which is the first quantitative survey of trypanosomatid shape of this magnitude. Data collection was focused on motile life cycle stages and identified examples of Trypanosoma in the bloodstream and invertebrate hosts, and Blastocrithidia, Leishmania, Phytomonas, Leptomonas, Crithidia, Herpetomonas, Strigomonas, Angomonas and Paratrypanosoma in invertebrate hosts. This represents coverage of the Trypanosoma, Phytomonas, Leishmaniinae, Blastocrithidia, Herpetomonas and the endosymbiont-bearing clades [25] and the newly-identified Paratrypanosoma genus which is the most basal known trypanosomatid lineage [26]. Descriptions of Leishmania, Phytomonas and Trypanosoma morphology in the invertebrate host were comparatively rare, reports were dominated by those of amastigotes from vertebrates, promastigotes from plants and trypomastigotes from vertebrates respectively.

For analysis these morphometric data required placement into morphological classes. There are well established morphological classes (trypomastigote, epimastigote, promastigote, choanomastigote, opisthomastigote and amastigote) for trypanosomatids which have historically been used to define the genera (Figure 1) [24], [27], [28]. Several of these genera have since been shown to be paraphyletic [25] indicating this degree of morphological subclassification is taxonomically deceptive. We therefore aimed to superclassify morphologies in a more biological relevant way guided by the phylogeny of trypanosomatids and the morphological transitions they can undergo through the life cycle. A comprehensive analysis of trypanosomatid morphological class occurrence by phylogeny would have been desirable, however there is little overlap between species description by morphology and by genetic data. Therefore we instead focused on fewer high quality descriptions of species morphology through the whole life cycle where both small subunit (SSU) rRNA and glycosomal glyceraldehydephosphate dehydrogenase (gGAPDH) sequence data (the most commonly sequenced genes for species identification and phylogenetic analysis of trypanosomatids) were available in GenBank [29]. This analysis revealed two distinct classes of life cycle: those which transition between trypomastigotes, epimastigotes and/or amastigotes, and those which transition between promastigotes, choanomastigotes, opisthomastigote and/or amastigotes (Figure 2). These life cycle patterns clustered by both SSU and gGAPDH phylogeny (Figure 2A and D). On this basis we defined two morphological superclasses which we tentatively named for the apparent morphological distinction of whether the trypanosomatid has an extended region of lateral flagellum attachment; ‘juxtaform’ (from the Latin juxta (beside), incorporating trypomastigotes and epimastigotes), or a free flagellum with no lateral attachment extending beyond the flagellar pocket neck region no lateral attachment; ‘liberform’ (from the Latin liber (free), incorporating promastigotes, choanomastigotes and opisthomastigotes).

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Figure 2. Trypanosomatid morphology, life cycle and phylogeny are indicative of two morphological superclasses.

A. Phylogeny of 12 representative trypanosomatids inferred from the small subunit (SSU) rRNA gene sequence, rooted with the outgroup B. saltans. Values at nodes indicate bootstrap support. The apparent paraphyly of Trypanosoma is a well documented example of a long branch attraction artefact [212]. B. Morphological classes attained though the 12 trypanosomatid life cycles. a[3], b[1], c[8], d[6], e[9], f[5], g[7], h[213], i[36], j[195], k[214], l[215], m[216], n[217], o[187], p[218], r[219], s[213]. C. Life cycle type and transmission route from the insect host in the 12 trypanosomatid life cycles. Relevance of the L. tarentolae amastigote in the life cycle [220], [221] and the transmission pathway [213] are debated. D. Phylogeny of the 12 trypanosomatids inferred from the glycosomal glyceraldehyde-3-phosphate dehydrogenase (gGAPDH), rooted with the outgroup B. saltans. Values at nodes indicate bootstrap support.

https://doi.org/10.1371/journal.pone.0079581.g002

The differences in morphology of juxtaforms and liberforms suggests there may be significant differences in the presence or molecular composition of major cytoskeletal components, particularly the flagellum attachment zone (FAZ), but also the paraflagellar rod (PFR), sub-pellicular microtubules, the flagellar pocket collar and the bilobe structure. However, a survey of presence or absence of homologs to known proteins in these structures in two representative juxtaforms (T. brucei and T. cruzi) and two representative liberforms (Leishmania mexicana and Crithidia fasciculata) with published genomes [30][33] did not reveal any clear groups of absent homologs (Figure 3). Therefore, in the absence of clear molecular markers for these two morphological superclasses, species were assigned to each superclass on the basis of genera for analysis; Trypanosoma and Blastocrithidia being juxtaform and Leishmania, Phytomonas, Leptomonas, Crithidia, Herpetomonas, Strigomonas, Angomonas and Paratrypanosoma being liberform. Genera were taken as those listed by Sergei Podlipaev [23], or any later reclassification. We identified one species with an ambiguous superclass arising from reclassification of its genus over time; Strigomonas culicis, which was previously classified as a Blastocrithidia [34]. Unlike the other Strigomonas species, S. culcis was analysed with the juxtaforms because of its epimastigote morphology.

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Figure 3. Conservation of known cytoskeletal-associated proteins between liberforms and juxtaforms.

Summary of cytoskeleton-associated proteins, the organism in which they were originally identified (red), and possession (black) or absence (white) of a homolog, identified by reciprocal best BLASTp. PFR1 and 2 were originally identified in Euglena gracilis. a[222], b[223], c[4], d[224], e[225], f[226], g[227], h[228], i[229], j[230], k[231], l[232], m[233], n[234], o[235], p[236], r[237], s[238], t[239].

https://doi.org/10.1371/journal.pone.0079581.g003

We analysed trypanosomatid shape and size diversity to determine whether limits to cell shape diversity across many trypanosomatid species correlate with different morphological classes, and what this implies for attainable trypanosomatid morphologies and their morphogenesis. Trypanosoma species have life cycle stages with juxtaform morphology both in and outside of the bloodstream where they may be subject to similar morphogenetic constraints, but different selective pressures. We therefore analysed the morphometric data particularly considering Trypanosoma species and whether there were limits to morphological diversity in different hosts which could be attributed to host selective pressures.

Morphological Diversity of Juxtaform Trypanosomatids

Our survey of trypanosomatid morphology from the bloodstream (see Materials and Methods) identified 110 isolates including examples from all vertebrate classes except reptiles and jawless fish. All isolates were Trypanosoma trypomastigotes. Trypanosomatid size and shape varied considerably; cell body length and width, free flagellum length and flagellum length each varied by at least a factor of 10 across species (Figure 4A to D, dark blue data series). Cell body width showed the greatest diversity (Figure 4D) and cell body length and flagellum length showed a similar distribution (Figure 4A and B). The smallest trypomastigotes were around 10 µm in length (Figure 4A), twice the typical maximum length of amastigotes of L. mexicana, T. cruzi and Leptomonas oncopelti [6], [35][37]. In order to assess if there were particular limits on diversity of trypomastigote cell shape, the correlations of these morphological features were analysed. In general there were only very weak correlations in parwise comparisons between free flagellum length, cell body width, cell body width and flagellum length for trypomastigotes (Figure 4E to J). Exceptions to this were cell body length and width, which showed a loose positive correlation (Figure 4E), and cell body length and flagellum length, which showed a strong positive correlation (Figure 4J). Linear regression for each plot indicated a significantly non-zero gradient (linear regression gradient test, 5% significance), indicating a weak tendency for larger trypanosomes to be larger in every respect. The lack of strong correlations of any morphological measure, with the exception that flagellum length appears to be linked with cell body length, indicates that the range of cell shapes that can be attained is large.

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Figure 4. Diversity and correlation of morphological features in species with a juxtaform morphology.

Data are colour coded by genus and host, Trypanosoma isolates are from either the vertebrate bloodstream of the insect host and all Blastocrithidia and Strigomonas isolates are from the insect host. n numbers for each plot are indicated in the top left. Solid black lines indicate the linear regression fit line for bloodstream Trypanosoma; the regression coefficient (R2) is shown each plot. Axenic procyclic form T. brucei dimensions [39], [41] and bloodstream form dimensions are indicated with a open data point.

https://doi.org/10.1371/journal.pone.0079581.g004

Insect-inhabiting juxtaform species were all members of either the Trypanosoma or Blastocrithidia genus. Our survey of trypanosomatids with a laterally attached flagellum from insect hosts yielded comparatively few (34) quantitative descriptions of morphology, and were dominated by epimastigotes of Blastocrithidia (25) and Strigomonas (1) species. Despite the large number of known Trypanosoma species which infect terrestrial animals, all expected to have a life cycle stage in an insect host, few morphological descriptions of Trypanosoma met our inclusion criteria. Trypanosoma are therefore underrepresented in this data set. Kinetoplast-posterior distance was not reported in any case which prevented analysis of total flagellum length. We analysed the morphology of these isolates to determine whether the diversity of trypomastigote morphologies seen in the bloodstream could also occur in juxtaform morphologies outside the bloodstream environment.

The range of cell body length, cell body width and free flagellum lengths reported in juxtaform isolates from the insect host were large (Figure 4A to D, light blue, brown and green data series), with a range approaching that of bloodstream-inhabiting trypomastigotes. The distribution of cell body length and width were significantly skewed towards smaller dimensions relative to trypomastigotes in the bloodstream (KS test, 5% significance) while there was no significant difference in the distribution of free flagellum length (KS test, 5% significance). Correlative analysis to assess morphological diversity in detail was not possible due to the small number of isolates, however the dimensions of parasites within these isolates was similar to that of small trypomastigotes (Figure 4E to J).

Morphological Diversity of Liberform Trypanosomatids

Many trypanosomatids inhabit invertebrate hosts, often within the alimentary tract and associated organs like the salivary glands. Promastigotes, choanomastigotes and opisthomastigotes (liberforms) are common in these environments, however epimastigotes and trypomastigotes (juxtaforms) and amastigotes are also widely reported. Our analysis of liberform morphology in the invertebrate host yielded quantitative morphological data from 103 isolates (see Materials and Methods), all of which were from insect hosts. Morphology diversity within these isolates was analysed to determine which morphological trends and limits observed within liberforms occur in juxtaforms, and vice versa.

Our survey of liberforms from insect hosts showed that both cell body and flagellum length vary by a factor of 10 (Figure 5A and B), a similar range to juxtaforms (Figure 4A to C). Cell width, however, only varied by a factor of approximately 3, with a maximum width of 4.5 µm reported (Figure 5C). This was unlike the large diversity of width of both trypomastigotes in the bloodstream and epimastigotes in the insect host (Figure 4D). The smallest liberforms, with several around 4 µm in length, were similar in size to amastigotes. Within the liberform superclass there was no significant difference between the distribution of cell body length, width and flagellum of the best-represented genera, Herpetomonas and Leptomonas (KS test, 5% significance). In contrast Crithidia, the third-best represented genus, had significantly shorter and wider cell bodies, and a shorter flagellum (KS test, 5% significance). The overall distribution of cell body length and width in liberforms were significantly skewed towards smaller dimensions relative to trypomastigotes in the bloodstream (KS test, 5% significance). The morphology of the early-branching Paratrypanosoma confusum [26] was not notable among the liberforms; all measured parameters were in the middle of the range (Figure 5A–F).

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Figure 5. Diversity and correlation of morphological features in species with a liberform morphology.

Data are colour coded by genus, all isolates are derived from the insect host. n numbers for each plot are indicated in the top left. Solid black lines indicate the linear regression fit line of data for all genera combined; the regression coefficient (R2) is shown each plot. Axenic promastigote L. mexicana dimensions [38] are indicated with an open data point.

https://doi.org/10.1371/journal.pone.0079581.g005

Limitations on cell shape diversity for liberforms were analysed through correlation of morphological measures. There was no strong correlation of cell body length, flagellum length or cell body width (Figure 5D to F). Linear regression for each plot indicated a significantly non-zero gradient (linear regression gradient test, 5% significance) only for the correlation of cell body length with free flagellum length (Figure 5F). This differed from cell body width in trypomastigotes in the bloodstream, which showed large diversity (Figure 4D) and a weak correlation with other morphological measures (Figure 4E to G). There therefore appears to be an additional constraint for a narrow cell width of liberforms in the invertebrate host, maintained across trypanosomatids inhabiting the full range of insect hosts. Of the 15 epimastigote isolates from invertebrates with width data available, 27% had widths greater than 4.5 µm (Figure 4D); this is wider than the largest width reported for liberforms. This suggests the constraint of liberform cells to a narrow range of widths may be intrinsic to cell organisation within this superclass, as juxtaforms (specifically epimastigotes) in invertebrate gut environments do not experience the same constraint. This limit on width is unlikely to be the only limit to morphological diversity, other selection-associated morphological adaptations associated with particular host properties are also likely.

Morphological Diversity within Isolates

In addition to recording the average morphological properties of isolates, the range of morphological variation within an isolate was also recorded for our meta-analysis. Within any individual isolate some degree of morphological variation arising from different life cycle and cell cycle stages would be anticipated, and this diversity provides a means to gain insight into the processes that may be involved.

We quantified the range of cell body and flagellum length variation by determining the ratio of the range to the mean, a dimensionless value we call the ‘length range ratio’. The cell body length range ratio varied greatly between isolates, ranging from near 0.0 (no range in length) to over 1.0 (Figure 6). The average cell body length range ratio in liberforms in the insect host was 0.74±0.51, significantly larger (Student’s t-test, 5% significance level) and with a wider distribution than trypomastigotes in both the insect and the bloodstream, 0.41±0.19 (Figure 6A and C). The mean length range ratio in these two classes were consistent with those that may be expected from the cell cycle, based on axenic promastigote L. mexicana where the length range ratio is 0.67 [38] and trypomastigote procyclic T. brucei where the length range ratio is 0.35 [39], [40]. The average cell body length range ratio of epimastigotes (Blastocrithidia species) was 0.73±0.37, similar to the liberforms and significantly larger (Student’s t-test, 5% significance) than trypomastigotes in the bloodstream (Figure 6A). Whether a large range in cell body length is associated with the host environment (for example reflecting more life cycle substages within the insect host), or is associated with an intrinsic aspect of cell organisation cannot be determined without more extensive data from trypomastigotes in the insect environment.

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Figure 6. Range of cell body and flagellum length within isolates of juxtaforms and liberforms.

Histograms of the ratio of the cell body length range within an isolate to the mean (the cell body ‘length range ratio’) (A and C) and flagellum length range ratio (B and D) for juxtaforms (A and B) and liberforms (C and D). n numbers for each plot are indicated in the bottom right. Vertical dashed line indicates where either proliferating axenic procyclic T. brucei trypomastigotes [39], [40] or L. mexicana promastigotes [38] would be positioned on the cell body length plots.

https://doi.org/10.1371/journal.pone.0079581.g006

Flagellum length showed a similar diversity to cell body length within isolates, with a mean flagellum length range ratio of 0.90±0.75 in liberforms in the insect host, significantly larger (Student’s t-test, 5% significance level) and with a wider distribution than trypomastigotes in the bloodstream, 0.51±0.25 (Figure 6B and D). This is consistent with the observed correlation of flagellum length with cell body length in trypomastigotes (Figure 4J), but not liberforms (Figure 5F), and the possible link of flagellum length with cell body length in juxtaforms. This is broadly consistent with the ranges of flagellum length which occur during axenic promastigote L. mexicana growth, in which a wide range of flagellar lengths are observed [38], and trypomastigote procyclic T. brucei division, where the flagellum length is less variable [41].

Extrinsic Limits on Trypomastigote Diversity in the Bloodstream

Having analysed whether there are limits on trypanosomatid morphological diversity which seem associated with the liberform or juxtaform morphological superclasses (presence or absence of a laterally attached flagellum), we can now address whether additional limits on cell shape occur associated with selective pressures. Selective pressures on trypanosomatid morphology would be expected to manifest as limits on shape diversity which do not correlate with morphological class, but instead with the host environment and its properties.

In this context Trypanosoma is notable as the only genus which inhabits the vertebrate bloodstream; it is also the only genus in which the trypomastigote morphology occurs. Furthermore it is the only genus defined by a unique morphological class, the trypomastigote, where the genus has been shown to be monophyletic by modern molecular phylogeny [25]. While Trypanosoma species can attain an epimastigote or amastigote morphology at some life cycle stages, they are almost universally found as a trypomastigote in the bloodstream environment. Trypanosoma lewisi is an exception to this where epimastigotes can be found in peripheral tissues, however even in this case parasites in the blood during high parasitaemias are trypomastigote [42]. This strong association of trypomastigote morphology with life in the bloodstream suggests there may also be associated selection pressures on trypomastigote size and shape; for example it has been suggested that T. brucei is of the correct size to allow effective motility amongst human erythrocytes [43]. The physical properties of a bloodstream environment are linked with the dimensions of host erythrocytes [44], therefore we aimed to detect selective pressures on trypomastigotes dimensions as limits on trypomastigote size diversity leading to a correlation of trypomastigote dimensions with host erythrocyte dimensions. Erythrocytes size is described by the dimensions of the major and minor axis of the elliptical face (Figure 7A) and, in the host species surveyed, these varied by a factor of 10 (Figure 7B), similar to the range of trypomastigote dimensions.

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Figure 7. Diversity of erythrocyte dimensions of hosts from which bloodstream Trypanosoma isolates were derived.

A. Diagram of erythrocyte measurements used to characterise their shape. B. Correlation of erythrocyte major axis and minor axis from vertebrate hosts of Trypanosoma parasites, subcategorised by class. n numbers for each genus are indicated in the bottom left. Homo sapiens erythrocyte dimensions are indicated with an open data point.

https://doi.org/10.1371/journal.pone.0079581.g007

Host erythrocyte major and minor axis did not show a clear correlation with trypanosomatid cell body length, width or free flagellum length (Figure 8). While there was little direct correlation other trends emerged: 95% of trypomastigotes were narrower than the host erythrocyte major axis (Figure 8C). Even large mammalian-infective Trypanosoma species, such as members of the monophyletic subgenus Megatrypanum, T. (M.) cervi and T. (M.) theileri [45], adhered to this width constraint. Using the test criterion of the number of trypanosomatids with a width smaller than the host erythrocyte major axis, a Monte Carlo permutation test indicated this trend was significant (5% significance level). 98% of trypomastigotes were also longer than the host erythrocyte major axis (Figure 8A) and all are longer than the minor axis (Figure 8B). Flagellum length, which correlates with cell body length (Figure 4J), showed a similar trend (Figure 8G and H). These two trends were, however, not significant by a Monte Carlo permutation test (5% significance level). Lack of significance likely arises from the large minimum trypomastigote cell length of approximately 10 µm.

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Figure 8. Limits on bloodstream-inhabiting trypanosomatid morphology relative to host erythrocyte dimensions.

Correlations of trypanosomatid cell body length, cell body width, free flagellum length and flagellum length with host erythrocyte major axis and minor axis. Solid data points indicate host species-specific erythrocyte data, where available, open data points indicate the average of erythrocyte data available for the host genus. Solid lines indicate the linear regression fit line for genus data; the regression coefficient (R2) is shown on each plot. Dashed lines indicate the line along which trypanosomatid and host erythrocyte dimensions would be equal. T. binneyi data points are circled with a dotted line. n numbers for each plot are indicated in the bottom right.

https://doi.org/10.1371/journal.pone.0079581.g008

One species which deviated from the apparent constraint on width by a large degree was identified. Trypanosoma binneyi has a large trypomastigote bloodstream form (47 to 67 µm long, 11 to 15 µm wide [46]) and can inhabit the bloodstream of the platypus (Ornithorhynchus anatinus) which has small erythrocytes similar to other mammals (5.2 µm diameter [47]). T. binneyi is a member of the fish and amphibian Trypanosoma clade [48], [49] and the platypus inhabits an aquatic environment. There is therefore the possibility that the platypus is not a normal host or is not the major host of this species, or that the platypus has only recently become a transmissive host of T. binneyi.

The universality of the trypomastigote (as opposed to epimastigote) morphology in the blood also supports the idea that a short kinetoplast posterior distance may be an adaptation for the bloodstream. As the nucleus is typically positioned near the centre of a trypomastigote the kinetoplast would be expected to be in the posterior half of the cell, however histograms of kinetoplast distance from the posterior (Figure 9A) and kinetoplast distance from the posterior as a fraction of cell body length (Figure 9B) for Trypanosoma spp. revealed no trend for kinetoplast/flagellar pocket positioning particularly close to the posterior of trypomastigotes in the bloodstream. Few trypomastigotes had a kinetoplast very close to the posterior, as observed in bloodstream form T. brucei, and many species had a kinetoplast positioned far from the posterior, often further than for procyclic form T. brucei. No particular bias of kinetoplast position between different host classes could be identified. While a short posterior–kinetoplast distance was not a feature of all trypomastigotes in the bloodstream, the kinetoplast is positioned less than 2 µm (≤10% cell body length) from the posterior in the bloodstream forms of several mammalian-infective African trypanosome species [50][57] (Figure 9C and D). Based on molecular phylogeny of 4 of these species (T. brucei, T. vivax, T. simiae and T. congolense), and the status of T. evansi as a petite mutant of T. brucei [58], these are members of the monophyletic T. brucei (Salivarian) clade [59], [60].

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Figure 9. Limits on kinetoplast position in bloodstream-inhabiting trypanosomatid species.

Histograms of kinetoplast-posterior (KP) distance (A and C) and the ratio of KP distance to cell body length (B and D) for all trypanosomatid isolates from the bloodstream (A and B) and a separate set of only mammalian-infective African trypanosomes (C and D). n numbers for each plot are indicated in the top right. Vertical dashed lines indicate where axenic PCF and BSF T. brucei would be positioned on these plots.

https://doi.org/10.1371/journal.pone.0079581.g009

Discussion

We have performed a meta-analysis of the morphologies of motile life cycle stages of a wide range of trypanosomatid parasites derived from two major environments they inhabit; the invertebrate gut and the vertebrate bloodstream. For analysis these were placed into two morphological superclasses, the liberforms and juxtaforms, defined from genera which can attain the promastigote, choanamastigote, opisthomastigote or amastigote morphology (Leishmania, Phytomonas, Herpetomonas, Leptomonas, Crithidia, Angomonas, Strigomonas and Paratrypanosoma) and the trypomastigote, epimastigote or amastigote morphology (Trypanosoma and Blastocrithidia) respectively. Strigomonas was the only genus where current classification indicates both juxtaforms (with one example, S. culcis) and liberforms (other species) occur [26]. We propose that these two morphological superclasses represent biologically meaningful groups. The fact that we could find no species which has the capacity to transition between morphologies unique to each of these superclasses through life cycle differentiation events (Figure 2B) supports this view. This was further supported by the clustering of these life cycle patterns by phylogeny inferred from SSU rRNA and gGAPDH gene sequences (Figure 2A and D), although juxtaform monoxenous insect trypanosomatids (Blastocrithidia and a subset of Strigomonas species) are currently poorly sampled.

Limits on diversity were analysed through the morphological parameters of cell body length and width, free and total flagellum length and kinetoplast position. Morphology of internal structures, such as the kinetoplast, were not considered in this analysis. The morphological parameters we analysed typically varied over a 10-fold range (Figure 4A to D and 5A to C). Within this range several patterns in morphological diversity of the trypanosomatids in different environments were identified. Firstly, juxtaforms and liberforms had different limits on cell shape diversity: the flagellum length of trypomastigotes, whilst in the bloodstream, was limited to a similar length as the cell body (Figure 4J) while in liberforms, whilst in an insect host, this was not the case (Figure 5G). Conversely, the cell body width in liberforms was limited to a narrow range (Figure 5D and E) while in juxtaforms it was not. In fact in trypomastigotes there was a loose positive correlation with cell size (Figure 2E). Juxtaforms were also limited to a larger minimum cell length, approximately 10 µm, (Figure 4A) than liberforms (Figure 5A).

Of the traditional kinetoplastid morphological classes (Figure 1) the trypomastigote is a notable juxtaform as it occurs near universally (we identified one exception, T. lewisi [42]) in vertebrate bloodstream-inhabiting life cycle stages, representing a near perfect coincidence of form and environment. Trypomastigotes in the bloodstream showed a trend for widths narrower than the minor axis of the host erythrocytes (Figure 8C). They also had lengths longer than the major axis of the host erythrocytes (Figure 8A), which may be associated with the limit on minimum cell length in morphologies with an attached flagellum (Figure 4A). Finally the flagellar pocket of the Salivarian clade of African mammalian-infective trypanosomes was located close to the posterior end of the cell, although in more diverse trypomastigotes this was not consistently observed.

We suggest these limits on morphology diversity can be explained through three groups of biological features; the physical properties of the host or vector environment, the mechanisms of cell morphogenesis through the cell cycle and, more speculatively, some functions of the flagellum in the host. The second of these is an intrinsic constraint of cell organisation, while the first and third represent extrinsic selective pressures from a host.

Extrinsic Morphological Constraints from the Host

Trypanosomatids need to fit through the capillaries of the host, the diameters of which average 80% of the largest erythrocyte dimension for vertebrate species [44]. The high mechanical rigidity of microtubules [61] in the sub-pellicular array would be expected to limit the degree to which trypanosomatid cells can deform to pass through the capillaries, restricting the range of morphologies which would be able to do so. The limited range of trypanosomatid widths (Figure 8C) is consistent with the idea that this mechanical effect exerts a selective pressure. It has previously been shown that the flagellum wavelength, cell body length and rotation of T. brucei while swimming confers a mechanical advantage when swimming amongst objects the size of human erythrocytes [43]. It is possible that the limit on minimum trypomastigote cell length (Figure 4A and 8G and H) indicates that this mechanism provides a similar advantage for motility of other species in their respective vertebrate hosts, although juxtaforms in the insect host shared this limit on minimum cell length.

Blood flow velocity in mammalian capillaries [62] is at least one order of magnitude higher than maximum T. brucei bloodstream form swimming speed [63], [64] making the role of swimming unclear as it is too slow to overcome the bulk flow of the blood. Nonetheless bloodstream form trypanosomes are typically highly active swimmers, and this has been confirmed in detail in vivo [65]. The function of this motility is debated, and there are many hypotheses including a role in tissue penetration (such as establishment of neurological stages of African sleeping sickness) or cell penetration (as for establishment of intracellular life cycle stages of Trypanosoma cruzi [66]). A moving cell body may also help escape from low affinity interaction with phagocytes, and avoid uptake. These hypotheses do not entail any particular requirement for a trypomastigote morphology, or any other specific morphological feature, for their function, and could therefore not be tested by analysing the observed range of trypomastigote morphologies.

In contrast, the proposed function of hydrodynamic surface clearance does suggest a particular advantageous morphology. It has been shown that hydrodynamic flow from swimming assists clearance of surface bound antibodies, and may assist avoidance of the adaptive immune response [67], [68]. Hydrodynamic flow forces arising from swimming concentrates large surface molecules, including bound immunoglobulins, at the posterior end of the cell. As endocytosis occurs exclusively within the flagellar pocket and trypanosomatids swim in the flagellum-first direction this suggests a trypomastigote morphology and a flagellar pocket located close to the posterior end would confer a selective advantage through an improved clearance rate [67], [68]. Our analysis indicated that the kinetoplast is consistently positioned near the posterior end in the T. brucei (Salivarian) clade (Figure 9). Members of this clade have a thick surface coat in the bloodstream [69], [70], and undergo antigenic variation via their monotypic coat of GPI-anchored variable surface glycoproteins (VSG) [71], [72]. The T. brucei VSG coat is internalised and recycled by an exceptionally high rate of endocytosis [73] and recycling of VSG with bound immunoglobulins is assisted by hydrodynamic effects [68]. The selective pressure would therefore be reduced antigenicity of the monotypic surface coat. While it appears this mechanism may have applied a selective pressure in the Salivarian clade it is not a general phenomenon amongst trypomastigotes; little constraint on kinetoplast location was identified in other species (Figure 9) so other methods of immune evasion may be dominant in those cases.

Intrinsic Morphological Constraints Associated with Morphogenesis

There are differences in the mechanisms of morphogenesis of different trypanosomatid morphologies, particularly in the regulation of cell body length, which may give rise to differing constraints on parasite morphology. In trypomastigote T. brucei, cell body length depends on flagellum length. Mutations which affect growth or attachment of the flagellum often give rise to abnormally short daughter cells [39], [74][76], and this mechanism appears to be used to cause morphological change during life cycle progression [77]. In contrast promastigote L. mexicana mutants with elongated or shortened flagella do not experience any large change to cell length or problems with division [78][80]. The differing degrees of correlation of cell body and flagellum length of juxtaforms and liberforms (Figure 4 and 5) suggest the morphogenesis of T. brucei and L. mexicana are representative of these two superclasses respectively, with lateral flagellum attachment and flagellum length mediating control of cell length in juxtaforms but not in liberforms. It seems plausible the greater minimum cell length observed in trypomastigotes is a morphogenetic constraint associated with this mechanism of cell length control, however further analysis of length regulation and morphogenesis in epimastigotes through the cell cycle will be required to confirm these hypotheses.

Unlike flagellum and cell body length, cell body width in liberforms seemed subject to an intrinsic constraint, limiting cells to a narrow range of widths (Figure 5), a constraint not seen in juxtaforms (Figure 4). This limit to morphological diversity was particularly striking and may be associated with the mechanisms used for cell body growth. Cell body morphogenesis in trypanosomatids is linked with the growth of the sub-pellicular microtubule array, which maintains a uniform coverage of equally spaced microtubules under the plasma membrane at all stages of the cell and life cycles. At least two mechanisms of array growth appear to occur, based on the identification of non-detyrosinated tubulin of newly polymerised microtubules in T. brucei [81]. The array can increase in length by polymerisation of a group of microtubules in a coordinated manner; this occurs at the posterior end of T. brucei and L. mexicana may undergo a similar growth at the posterior end [38], [82]. New microtubules can also be inserted into the existing array, leading to an increased width of that section of the cytoskeleton; this is thought to occur throughout the mid portion of the cell body during later stages of division in T. brucei [81] (Figure 10B). Steady growth in width is not a morphogenetic process used by L. mexicana, which instead undergoes a rapid remodelling in shape with an increase in width and decrease in length in the lead up to cytokinesis [38] (Figure 10A).

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Figure 10. Cell body morphogenesis of a promastigote and trypomastigote through the cell cycle.

Cell growth is indicated by arrows with a perpendicular bar, the bar indicates the region that grows. Cell remodelling is indicated by barb-headed arrows. The path of furrow ingression is indicated by dashed arrows. The old and new flagellum are coloured red and turquoise respectively. A. Summary of the morphological changes of an example liberform (promastigote L. mexicana) through the cell cycle [38]. The cell grows in length, then remodels the cytoskeleton during cytokinesis. B. Summary of the morphological changes of an example juxtaform (trypomastigote procyclic T. brucei) through the cell cycle [39], [81], [240]. The cell first grows in length then in width prior to cytokinesis.

https://doi.org/10.1371/journal.pone.0079581.g010

This quantitative analysis of mechanisms used for morphogenesis of T. brucei procyclic trypomastigotes and L. mexicana promastigotes (Figure 10), combined with the apparent constraint on cell widths attained by liberforms (Figure 5D and E), may indicate it is a general phenomenon that liberforms use rapid cellular remodelling instead of gradual growth for morphogenesis; it may be the case that the second method of array growth, increase in width through microtubule insertion, does not occur in this superclass. Conversely the lateral attachment of the flagellum in trypomastigotes may prevent the dramatic rearrangement of cell shape used by Leishmania for cytokinesis and enforce a different mechanism for increasing in width. Again analysis of the morphogenesis of epimastigotes through the cell cycle will be required to test if this is a universal feature of juxtaform morphogenesis. Existing qualitative descriptions and diagrams of dividing liberform trypanosomatids, particularly Leptomonas spp. [83], [84], and juxtaform trypanosomatids, particularly mammalian-infective trypanosomes [22] also support this hypothesis. Furthermore the range of cell body and flagellum lengths seen within isolates is broadly consistent with morphological variation as part of a cell cycle similar to L. mexicana or T. brucei for liberforms and juxtaforms respectively (Figure 6). Together, and in combination with apparent difference in role of the flagellum in cell body length regulation, these further support the concept of the liberform and juxtaform morphological superclasses and that differences in their morphogenesis may impose different constraints on cell shape.

There has been growing evidence that definition of trypanosomatid genera by the morphological classes they attain in the life cycle does not reflect the molecular phylogeny of these species, and there have since been efforts to regroup the species into monophyletic subfamilies on the basis of genetic analyses [25]. The recently-identified promastigote Paratrypanosoma confusum strongly suggests the ancestral morphology of trypanosomatids was liberform [26], implying the juxtaform morphology has arisen in at least one well-sampled lineage (Trypanosoma) and two less well-sampled lineages (Blastocrithida and Strigomonas). We suggest that morphological data still has biological value in this context; Trypanosoma are universally juxtaform and unique in their capacity to live in the vertebrate bloodstream, and the superclassification of trypanosomatid morphologies into the liberform and juxtaform superclasses correlates well with the ability for parasite morphological change through the life cycle (Figure 2) and capacity for morphological diversity (Figure 4 and 5), independent of whether juxtaforms or liberforms prove to be mono- or paraphyletic. Our analysis showed that protein components of cytoskeletal structures particularly associated with juxtaforms or liberforms respectively, have not been lost or gained in a modular fashion. The FAZ being the clearest example of this. Clearly identifiable homologues of the majority of genes encoding such proteins are present in members of the other superclass (Figure 3). This suggests that the mechanisms that underlie morphological differences are better interpreted as modulation of the cytoskeleton rather than modular loss or gain of the capacity to form particular cytoskeletal structures, like the FAZ. It may therefore be unsurprising that multiple lineages seem to have independently evolved a juxtaform morphology.

The amastigote morphology, which is poorly defined, occurs in proliferative intracellular Leishmania and Trypanosoma [10][12], encysted life cycle stages of monoxenous insect parasites [14] and occasionally other forms. It cannot be directly classified as juxtaform or liberform as it lacks a long external flagellum. One prediction of the superclass model is the possibility that juxtaform and liberform amastigotes may be heavily modulated examples of epimastigotes and promastigotes respectively; there may be morphological differences between the juxtaform and liberform amastigote in the structure of the flagellum exit from the pocket, the ultrastructure of the collar and neck of the pocket, and presence or absence of an elongated FAZ-like structure.

The Function of the Flagellum

The classic morphological classes and our proposed juxtaform and liberform superclasses are all defined on the basis of the flagellum, and as a result many of these discussion points touch aspects of flagellum function in trypanosomatids. It is valuable to consider their implications for the broader question of why does any trypanosomatid have a particular morphology at any particular life cycle stage, and what is the function of the flagellum within that morphology. The flagellum, which is universally present in trypanosomatids, is a central component in the vital flagellar pocket structure; it is the flagellum and its interaction with the flagellar pocket collar which defines the pocket structure and orientation [85]. Endocytosis (and the associated processes of receptor mediated uptake, surface recycling and antibody clearance) occur only in this membrane domain therefore it appears the minimal trypanosomatid cell requires at least a short flagellum to maintain this structure. This is consistent with the presence of very short flagella in amastigotes, whether living inside the cytoplasm (like T. cruzi), a parasitophorous vacuole (like Leishmania) or extracellularly (like the monoxenous insect trypanosomatids) [21], [86]. In addition to its role in the formation of the flagellar pocket, the trypanosomatid flagellum has at least four other major functions: surface attachment, motility, control of morphogenesis (as described above) and sensation (for which there is still limited evidence).

Surface attachment in the insect is common in most trypanosomatids [28] and is usually mediated via the flagellum. Attachment is associated with formation of hemidesmosome-like structures which have been described in detail in diverse genera; Crithidia [87], Leptomonas [88], Trypanosoma [89][93] and Leishmania [94][96]. These descriptions all concern epimastigotes or promastigotes. While the trypomastigote flagellum is capable of adhesion to a surface (its own cell body via the FAZ [97]) there are no known examples of trypomastigotes adhering to external surfaces, such as a fly intestine. In the non-trypomastigote morphologies, the ability to attach to surfaces may be a vital and limiting role of the extended free flagellum. Further analysis will be required to determine what constraints on morphology have arisen from selective pressures exerted by the invertebrate host.

The universality of the trypomastigote morphology and its motile flagellum in the bloodstream suggest a vital role of this morphology, which we speculate may be primarily associated with the motility of the flagellum. While trypomastigote life cycle stages do occur outside of the blood (for example the T. brucei procyclic (proliferative) and metacyclic (non-proliferative) forms [1], [98]) they are comparatively rare in the vector. Instead Trypanosoma spp. often have an epimastigote morphology, like Blastocrithidia, in the insect. It is therefore tempting to speculate that the trypomastigote morphology is a specialised juxtaform morphology, evolution of which was a key event in adaptation to the bloodstream environment by an ancestral monoxenous insect parasite. Within the bloodstream we have shown there are limits on kinetoplast position, cell body width and cell body and flagellum length which appear linked with plausible selective pressures associated with surface coat recycling (consistent with a motility role of the flagellum) and the size of host capillaries. Furthermore there appear to be particular constraints on cell morphogenesis, consistent with a morphogenetic role of the flagellum, associated specifically with the juxtaform cell shape. These findings give clear insights into functions of the typomastigote cell shape and its long flagellum. With the ability to generate morphological mutants in the laboratory and to compare genomes of diverse trypanosomatids, it is now feasible to search for the underlying principles that shape the trypanosomatid cell.

Materials and Methods

Literature concerning identification of new species and surveys of prevalence of trypanosomatid infections was systematically surveyed from the literature. Cell body length, cell body width, free flagellum length and kinetoplast to posterior distance were recorded, but morphological data was only included if certain criteria were met:

For bloodstream-inhabiting species, trypanosomatid dimensions from isolates were recorded if the parasite cell body length and width were available, host genus erythrocyte dimensions were available in the Erythrocyte Size Database [99] and if measurements were taken from blood isolates or axenic cultures immediately derived from isolates and if measurements were representative of the complete range of morphologies present. Erythrocyte major and minor axes were recorded for the host species (if available) in addition to the average erythrocyte major and minor axes across all species in the host genus. This generated a data set covering morphology of 110 isolates from the blood of 73 host vertebrates (covering 48 genera distributed across Amphibia, Aves, Chondrichthyes, Mammalia and Osteichthyes) (Table S1). Trypanosoma morphology references: [5], [47], [100][118], [118][140]. Original erythrocyte dimensions references: [47], [141][167].

Trypanosoma morphology data were supplemented with measurements of cell body length and kinetoplast-posterior distance from previously published illustrations (using ImageJ [168]) of the mammalian-infective African trypanosomes T. brucei, T. evansi, T. vivax, T. simoae, T. congolense, T. nanum, T. pecorum and T. uniforme. Original host species for these isolates was not clear, therefore these data were not included in the complete trypomastigote/erythrocyte morphology data set. Mammalian-infective African trypanosome references: [50][57].

For species inhabiting an invertebrate host, dimensions were recorded if parasite cell body and flagellum length were available, if measurements were taken from the initial isolate or axenic cultures immediately derived from isolates and if measurements were representative of the complete range of morphologies present. Only flagellated life cycle stages were included. This generated a data set covering morphology of 103 isolates of trypanosomatids with a free flagellum from 73 host insects (covering 63 genera) (Table S1) and a data set covering morphology of 35 isolates of trypanosomatids with a laterally attached flagellum from 27 host insects (covering 25 genera) (Table S1). Crithidia, Leptomonas, Herpetomonas, Leishmania, Phytomonas, Angomonas, Strigomonas and Paratrypanosoma references: [24], [26], [34], [83], [84], [169][201]. Trypanosoma and Blastocrithidia references: [24], [169], [170], [181], [182], [202], [203].

Mean, minimum, maximum and standard deviation of morphological measurements were recorded where available, where mean length was not provided it was derived from the mean of the minimum and maximum. Where minimum and maximum were not available they were estimated as the 5th and 95th percentiles (i.e. expected minimum and maximum for a sample of n = 20) from the mean and standard deviation where possible. Flagellum length of morphologies with a laterally attached flagellum was estimated from the sum of cell body length and free flagellum length, minus the kinetoplast-posterior distance. Minimum and maximum flagellum length were estimated using the root sum squared deviation of the minimum and maximum of cell body length, free flagellum length and kinetoplast-posterior distance from their respective means. All plots concerning trypanosomatid morphology are derived from these data and n numbers indicate the number of species for which the morphometric data required for the plot was available.

Trypanosomatid phylogeny was inferred from SSU and gGAPDH gene sequence data available in Genbank. Unless otherwise indicated sequences were identified by species name and when sequences from multiple clones or isolates were available a single representative sequence was selected by sequence length (complete gene sequences were preferable) and quality (no gaps or uncalled bases). Multiple sequence alignments were generated with ClustalW [204], Clustal Omega [205], Kalign [206], MAFFT [207], MUSCLE [208] and T-Coffee [209] then combined using MergeAlign [210] and the alignment trimmed to only columns with a MergeAlign score over 0.5. The phylogenetic tree was estimated by the neighbour joining method [211] with 1,000 bootstrap iterations. Nodes with less than 50% (0.50) bootstrap support were collapsed to polytomies, preserving branch length. Genbank accession numbers: SSU: M12676.1, FJ900241.2, AJ223566.1, AJ009140.1, AJ009143.1, GQ332360.1, X53913.1, M84225.1, U39577.1, Y00055.1, AF153037.2, L18872.1, DQ207573.1. gGAPDH: AJ620263.1, AF047493.1, DQ092548.1, EU084894.1, M26816.1, XM_001566870.1, XM_003877392.1, DQ092549.1, AJ620272.1, X52898.1, AJ620247.1, FJ968528.1, EU084900.1.

Presence and absence of homologs of proteins associated with cytoskeletal structures were identified by the criterion of reciprocal best BLASTp against the predicted protein sequences of a genome, with a minimum p-value cutoff of 10−5. Analysis was performed online using TriTrypDB.org [30].

Supporting Information

Table S1.

Morphometry of trypanosomatids. The complete morphometric data set used for this analysis, derived from previously published data for many trypanosomatid species from different vertebrate and invertebrate hosts.

https://doi.org/10.1371/journal.pone.0079581.s001

(XLSX)

Author Contributions

Conceived and designed the experiments: RJW. Analyzed the data: RJW EG KG. Wrote the paper: RJW EG KG. Contributed to the literature survey data collection: RJW EG KG.

References

  1. 1. Vickerman K (1985) Developmental Cycles and Biology of Pathogenic Trypanosomes. Br Med Bull 41: 105–114.
  2. 2. Vickerman K (1994) The evolutionary expansion of the trypanosomatid flagellates. Int J Parasitol 24: 1317–1331.
  3. 3. Hoare CA, Wallace FG (1966) Developmental Stages of Trypanosomatid Flagellates: a New Terminology. Nature 212: 1385–1386
  4. 4. De Jesus AR, Cooper R, Espinosa M, Gomes JE, Garcia ES, et al. (1993) Gene deletion suggests a role for Trypanosoma cruzi surface glycoprotein GP72 in the insect and mammalian stages of the life cycle. J Cell Sci 106 (Pt 4): 1023–1033.
  5. 5. Baker JR (1956) Studies on Trypanosoma avium Danilewsky 1885 III. Life Cycle in vertebrate and invertebrate hosts. Parasitology 46: 335–352
  6. 6. Goldenberg S, Avila AR (2011) Aspects of Trypanosoma cruzi stage differentiation. Adv Parasitol 75: 285–305
  7. 7. Letch CA (1980) The life-cycle of Trypanosoma cobitis Mitrophanow 1883. Parasitology 80: 163–169.
  8. 8. Matthews KR (2005) The developmental cell biology of Trypanosoma brucei. J Cell Sci 118: 283–290
  9. 9. Minchin E, Thompson J (1915) The Rat-Trypanosome, Trypanosoma lewisi, in its relation to the Rat-Flea, Ceratophyllus fasciatus. Quart J Microsc Sci n s 60: 463–962.
  10. 10. Engel JC, Dvorak JA (1988) Trypanosoma cruzi: Cell Biological Behavior of Epimastigote and Amastigote Forms in Axenic Culture. Journal of Eukaryotic Microbiology 35: 513–518
  11. 11. Bates PA (1994) Complete developmental cycle of Leishmania mexicana in axenic culture. Parasitology 108: 1–9.
  12. 12. Tomlinson S, Vandekerckhove F, Frevert U, Nussenzweig V (1995) The Induction of Trypanosoma Cruzi Trypomastigote to Amastigote Transformation by Low pH. Parasitology 110: 547–554
  13. 13. Angelopoulos E (1970) Pellicular microtubules in the family Trypanosomatidae. J Protozool 17: 39–51.
  14. 14. Romeiro A, Solé-Cava A, Sousa MA, De Souza W, Attias M (2000) Ultrastructural and Biochemical Characterization of Promastigote and Cystic Forms of Leptomonas wallacei n. sp. Isolated from the Intestine of its Natural Host Oncopeltus fasciatus (Hemiptera: Lygaeidae). Journal of Eukaryotic Microbiology 47: 208–220
  15. 15. Ogbadoyi EO, Robinson DR, Gull K (2003) A high-order trans-membrane structural linkage is responsible for mitochondrial genome positioning and segregation by flagellar basal bodies in trypanosomes. Mol Biol Cell 14: 1769–1779
  16. 16. Robinson DR, Gull K (1991) Basal body movements as a mechanism for mitochondrial genome segregation in the trypanosome cell cycle. Nature 352: 731–733
  17. 17. Green HP, Del Pilar Molina Portela M, St Jean EN, Lugli EB, Raper J (2003) Evidence for a Trypanosoma brucei lipoprotein scavenger receptor. J Biol Chem 278: 422–427
  18. 18. McConville MJ, Mullin KA, Ilgoutz SC, Teasdale RD (2002) Secretory pathway of trypanosomatid parasites. Microbiol Mol Biol Rev 66: 122–154; table of contents.
  19. 19. Morgan GW, Hall BS, Denny PW, Carrington M, Field MC (2002) The kinetoplastida endocytic apparatus. Part I: a dynamic system for nutrition and evasion of host defences. Trends Parasitol 18: 491–496.
  20. 20. Morgan GW, Hall BS, Denny PW, Field MC, Carrington M (2002) The endocytic apparatus of the kinetoplastida. Part II: machinery and components of the system. Trends Parasitol 18: 540–546.
  21. 21. Gull K (2003) Host–parasite interactions and trypanosome morphogenesis: a flagellar pocketful of goodies. Current Opinion in Microbiology 6: 365–370
  22. 22. Hoare CA (1972) The trypanosomes of mammals: a zoological monograph. Blackwell Scientific Publications. 808 p.
  23. 23. Podlipaev SA (1990) Catalogue of world fauna of Trypanosomatidae (Protozoa). Proceedings of the Zoological Institute of the USSR Academy of Sciences 217: 1–177.
  24. 24. Wallace FG (1966) The trypanosomatid parasites of insects and arachnids. Experimental Parasitology 18: 124–193
  25. 25. Maslov DA, Votýpka J, Yurchenko V, Lukeš J (2013) Diversity and phylogeny of insect trypanosomatids: all that is hidden shall be revealed. Trends Parasitol 29: 43–52
  26. 26. Flegontov P, Votýpka J, Skalický T, Logacheva MD, Penin AA, et al.. (2013) Paratrypanosoma Is a Novel Early-Branching Trypanosomatid. Curr Biol. doi:10.1016/j.cub.2013.07.045.
  27. 27. Hoare CA (1964) Morphological and Taxonomic Studies on Mammalian Trypanosomes. X. Revision of the Systematics. Journal of Eukaryotic Microbiology 11: 200–207
  28. 28. McGhee RB, Cosgrove WB (1980) Biology and physiology of the lower Trypanosomatidae. Microbiol Rev 44: 140–173.
  29. 29. Benson DA, Karsch-Mizrachi I, Lipman DJ, Ostell J, Wheeler DL (2005) GenBank. Nucleic Acids Res 33: D34–D38
  30. 30. Aslett M, Aurrecoechea C, Berriman M, Brestelli J, Brunk BP, et al. (2009) TriTrypDB: a functional genomic resource for the Trypanosomatidae. Nucleic Acids Research 38: D457–D462
  31. 31. Berriman M, Ghedin E, Hertz-Fowler C, Blandin G, Renauld H, et al. (2005) The genome of the African trypanosome Trypanosoma brucei. Science 309: 416–422
  32. 32. El-Sayed NM, Myler PJ, Bartholomeu DC, Nilsson D, Aggarwal G, et al. (2005) The genome sequence of Trypanosoma cruzi, etiologic agent of Chagas disease. Science 309: 409–415
  33. 33. Peacock CS, Seeger K, Harris D, Murphy L, Ruiz JC, et al. (2007) Comparative genomic analysis of three Leishmania species that cause diverse human disease. Nat Genet 39: 839–847
  34. 34. Teixeira MMG, Borghesan TC, Ferreira RC, Santos MA, Takata CSA, et al. (2011) Phylogenetic validation of the genera Angomonas and Strigomonas of trypanosomatids harboring bacterial endosymbionts with the description of new species of trypanosomatids and of proteobacterial symbionts. Protist 162: 503–524
  35. 35. Mcghee RB, Hanson WL (1964) Comparison of the Life Cycle of Leptomonas oncopelti and Phytomonas elmassiani. Journal of Eukaryotic Microbiology 11: 555–562
  36. 36. Handman E, Bullen DVR (2002) Interaction of Leishmania with the host macrophage. Trends Parasitol 18: 332–334.
  37. 37. Malysheva MN, Frolov AO, Skarlato SO (2006) [Development of cyst-like cells of the flagellate Leptomonas oncopelti in the midgut of the hemipteran Oncopeltus fasciatus]. Tsitologiia 48: 723–733.
  38. 38. Wheeler RJ, Gluenz E, Gull K (2011) The cell cycle of Leishmania: morphogenetic events and their implications for parasite biology. Mol Microbiol 79: 647–662
  39. 39. Robinson DR, Sherwin T, Ploubidou A, Byard EH, Gull K (1995) Microtubule polarity and dynamics in the control of organelle positioning, segregation, and cytokinesis in the trypanosome cell cycle. J Cell Biol 128: 1163–1172.
  40. 40. Wheeler RJ, Gull K, Gluenz E (2012) Detailed interrogation of trypanosome cell biology via differential organelle staining and automated image analysis. BMC Biology 10: 1
  41. 41. Farr H, Gull K (2009) Functional studies of an evolutionarily conserved, cytochrome b5 domain protein reveal a specific role in axonemal organisation and the general phenomenon of post-division axonemal growth in trypanosomes. Cell Motil Cytoskeleton 66: 24–35
  42. 42. Deane MP (1969) On the life cycle of trypanosomes of the lewisi group and their relationships to other mammalian trypanosomes. Rev Inst Med Trop São Paulo 11: 34–43.
  43. 43. Heddergott N, Krueger T, Wei A, Stellamans E, Uppaluri S, et al.. (2011) Trypanosome motion represents an adaptation to the environment of the vertebrate bloodstream. Proceedings of The Kinetoplastid Molecular Cell Biology Meeting. Woods Hole, MA, USA. Available: http://hermes.mbl.edu/kmcb/2011/kmcb_book_single.pdf. Accessed 13 June 2012.
  44. 44. Snyder GK, Sheafor BA (1999) Red Blood Cells: Centerpiece in the Evolution of the Vertebrate Circulatory System. Amer Zool 39: 189–198
  45. 45. Rodrigues AC, Paiva F, Campaner M, Stevens JR, Noyes HA, et al. (2006) Phylogeny of Trypanosoma (Megatrypanum ) theileri and related trypanosomes reveals lineages of isolates associated with artiodactyl hosts diverging on SSU and ITS ribosomal sequences. Parasitology 132: 215–224
  46. 46. Mackerras MJ (1959) The haematozoa of Australian mammals. Australian Journal of Zoology 7: 105–135.
  47. 47. Collins GH, Whittington RJ, Canfield PJ (1986) Theileria ornithorhynchi Mackerras, 1959 in the platypus, Ornithorhynchus anatinus (Shaw). J Wildlife Dis 22: 19–24.
  48. 48. Jakes KA, O’Donoghue PJ, Adlard RD (2001) Phylogenetic relationships of Trypanosoma chelodina and Trypanosoma binneyi from Australian tortoises and platypuses inferred from small subunit rRNA analyses. Parasitology 123: 483–487.
  49. 49. Gibson WC, Lom J, Peckova H, Ferris VR, Hamilton PB (2005) Phylogenetic analysis of freshwater fish trypanosomes from Europe using ssu rRNA gene sequences and random amplification of polymorphic DNA. Parasitology 130: 405–412.
  50. 50. Bruce D, Hamerton AE, Bateman HR (1909) A trypanosome from Zanzibar. Proc R Soc B 81: 14–30.
  51. 51. Bruce D, Hamerton AE, Bateman HR, Mackie FP (1910) Trypanosome diseases of domestic animals in Uganda. I.-Trypanosoma pecorum. Proc R Soc B 82: 468–479.
  52. 52. Bruce D, Hamerton AE, Bateman HR, Mackie FP (1910) Trypanosome diseases of domestic animals in Uganda. II:-Trypanosoma brucei (Plimmer and Bradford). Proc R Soc B 83: 1–14.
  53. 53. Bruce D, Hamerton AE, Bateman HR, Mackie FP (1910) Trypanosome diseases of domestic animals in Uganda. III.-Trypanosoma vivax (Ziemann). Proc R Soc B 83: 15–27.
  54. 54. Bruce D, Hamerton AE, Bateman HR, Mackie FP (1911) Trypanosome diseases of domestic animals in Uganda. IV.-Trypanosoma uniforme, sp. nov. Proc R Soc B 83: 176–179.
  55. 55. Bruce D, Hamerton AE, Bateman HR, Mackie FP (1911) Trypanosome diseases of domestic animals in Uganda. V.-Trypanosoma nanum (Laveran). Proc R Soc B 83: 180–186.
  56. 56. Bruce D, Harvey D, Hamerton AE, Davey JB, Bruce ME (1912) The morphology of Trypanosoma simiae, sp. nov. Proc R Soc B 85: 477–481.
  57. 57. Bruce D (1911) The morphology of Trypanosoma evansi (Steel). Proc R Soc B 84: 181–187.
  58. 58. Lai D-H, Hashimi H, Lun Z-R, Ayala FJ, Lukeš J (2008) Adaptations of Trypanosoma brucei to gradual loss of kinetoplast DNA: Trypanosoma equiperdum and Trypanosoma evansi are petite mutants of T. brucei. PNAS 105: 1999–2004
  59. 59. Stevens J, Gibson W (1999) The evolution of salivarian trypanosomes. Mem Inst Oswaldo Cruz 94: 225–228.
  60. 60. Hamilton PB, Stevens JR, Gaunt MW, Gidley J, Gibson WC (2004) Trypanosomes are monophyletic: evidence from genes for glyceraldehyde phosphate dehydrogenase and small subunit ribosomal RNA. Int J Parasitol 34: 1393–1404
  61. 61. Gittes F, Mickey B, Nettleton J, Howard J (1993) Flexural rigidity of microtubules and actin filaments measured from thermal fluctuations in shape. J Cell Biol 120: 923–934.
  62. 62. Ivanov KP, Kalinina MK, Levkovich YI (1981) Blood flow velocity in capillaries of brain and muscles and its physiological significance. Microvascular Research 22: 143–155
  63. 63. Uppaluri S, Nagler J, Stellamanns E, Heddergott N, Herminghaus S, et al. (2011) Impact of microscopic motility on the swimming behavior of parasites: straighter trypanosomes are more directional. PLoS Comput Biol 7: e1002058
  64. 64. Weiße S, Heddergott N, Heydt M, Pflästerer D, Maier T, et al. (2012) A Quantitative 3D Motility Analysis of Trypanosoma brucei by Use of Digital In-line Holographic Microscopy. PLoS ONE 7: e37296
  65. 65. Frevert U, Movila A, Nikolskaia OV, Raper J, Mackey ZB, et al. (2012) Early Invasion of Brain Parenchyma by African Trypanosomes. PLoS ONE 7: e43913
  66. 66. Fernandes MC, Cortez M, Flannery AR, Tam C, Mortara RA, et al. (2011) Trypanosoma cruzi subverts the sphingomyelinase-mediated plasma membrane repair pathway for cell invasion. J Exp Med 208: 909–921
  67. 67. Dean SD, Matthews KR (2007) Restless gossamers: antibody clearance by hydrodynamic flow forces generated at the surface of motile trypanosome parasites. Cell Host Microbe 2: 279–281
  68. 68. Engstler M, Pfohl T, Herminghaus S, Boshart M, Wiegertjes G, et al. (2007) Hydrodynamic flow-mediated protein sorting on the cell surface of trypanosomes. Cell 131: 505–515
  69. 69. Vickerman K (1969) On the surface coat and flagellar adhesion in trypanosomes. Journal of cell science 5: 163–193.
  70. 70. Vickerman K (1969) The Fine Structure of Trypanosoma congolense in Its Bloodstream Phase. Journal of Eukaryotic Microbiology 16: 54–69
  71. 71. Cross GAM (1990) Cellular and genetic aspects of antigenic variation in trypanosomes. Annual review of immunology 8: 83–110.
  72. 72. Jackson AP, Allison HC, Barry JD, Field MC, Hertz-Fowler C, et al.. (2013) A Cell-surface Phylome for African Trypanosomes. PLoS Negl Trop Dis 7. Available: http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3605285/. Accessed 9 April 2013.
  73. 73. Engstler M, Thilo L, Weise F, Grünfelder CG, Schwarz H, et al. (2004) Kinetics of endocytosis and recycling of the GPI-anchored variant surface glycoprotein in Trypanosoma brucei. J Cell Sci 117: 1105–1115
  74. 74. Ralston KS, Kabututu ZP, Melehani JH, Oberholzer M, Hill KL (2009) The Trypanosoma brucei flagellum: moving parasites in new directions. Annu Rev Microbiol 63: 335–362
  75. 75. Vaughan S (2010) Assembly of the flagellum and its role in cell morphogenesis in Trypanosoma brucei. Curr Opin Microbiol 13: 453–458
  76. 76. Vaughan S, Gull K (2008) The structural mechanics of cell division in Trypanosoma brucei. Biochem Soc Trans 36: 421–424
  77. 77. Sharma R, Peacock L, Gluenz E, Gull K, Gibson W, et al. (2008) Asymmetric cell division as a route to reduction in cell length and change in cell morphology in trypanosomes. Protist 159: 137–151
  78. 78. Wiese M, Kuhn D, Grünfelder CG (2003) Protein kinase involved in flagellar-length control. Eukaryotic Cell 2: 769–777.
  79. 79. Bengs F, Scholz A, Kuhn D, Wiese M (2005) LmxMPK9, a mitogen-activated protein kinase homologue affects flagellar length in Leishmania mexicana. Mol Microbiol 55: 1606–1615
  80. 80. Erdmann M, Scholz A, Melzer IM, Schmetz C, Wiese M (2006) Interacting protein kinases involved in the regulation of flagellar length. Mol Biol Cell 17: 2035–2045
  81. 81. Sherwin T, Gull K (1989) Visualization of detyrosination along single microtubules reveals novel mechanisms of assembly during cytoskeletal duplication in trypanosomes. Cell 57: 211–221.
  82. 82. Ambit A, Woods KL, Cull B, Coombs GH, Mottram JC (2011) Morphological events during the cell cycle of Leishmania major. Eukaryotic Cell. Available: http://www.ncbi.nlm.nih.gov/pubmed/21926331. Accessed 25 September 2011.
  83. 83. Vickerman K (1960) Herpetomonas ludwigi (Kramar, 1950) n. comb., the the trypanosomatid parasite of cranefly larvae (Diptera, Tipulidae). Pathol Biol 50: 351–363.
  84. 84. Wallace FG, Todd SR (1964) Leptomonas mirabilis Roubaud 1908 in a Central American Blowfly. Journal of Eukaryotic Microbiology 11: 502–505.
  85. 85. Absalon S, Blisnick T, Bonhivers M, Kohl L, Cayet N, et al. (2008) Flagellum elongation is required for correct structure, orientation and function of the flagellar pocket in Trypanosoma brucei. J Cell Sci 121: 3704–3716
  86. 86. Gull K (2009) The parasite point of view: insights and questions on the cell biology of Trypanosoma and Leishmania parasite-phagocyte interactions. In: Russell DG, Gordon S, editors. Phagocyte-Pathogen Interactions: Macrophages and the Host Response to Infection. Washington: ASM Press. 453–462.
  87. 87. Brooker BE (1971) Flagellar attachment and detachment of Crithidia fasciculata to the gut wall of Anopheles gambiae. Protoplasma 73: 191–202.
  88. 88. Laugé G, Nishioka RS (1977) Ultrastructural study of the relations between Leptomonas oncopelti (Noguchi and Tilden), protozoa trypanosomatidae, and the rectal wall of adults of Oncopeltus fasciatus dallas, hemiptera lygaeidae. Journal of Morphology 154: 291–305
  89. 89. Molyneux DH (1969) The fine-structure of the epimastigote forms of Trypanosoma lewisi in the rectum of the flea, Nosopsyllus fasciatus. Parasitology 59: 55–66.
  90. 90. Vickerman K (1973) The Mode of Attachment of Trypanosoma vivax in the Proboscis of the Tsetse Fly Glossina fuscipes: an Ultrastructural Study of the Epimastigote Stage of the Trypanosome. Journal of Eukaryotic Microbiology 20: 394–404
  91. 91. Thévenaz P, Hecker H (1980) Distribution and attachment of Trypanosoma (Nannomonas) congolense in the proximal part of the proboscis of Glossina morsitans morsitans. Acta Trop 37: 163–175.
  92. 92. Beattie P, Gull K (1997) Cytoskeletal architecture and components involved in the attachment of Trypanosoma congolense epimastigotes. Parasitology 115: 47–55.
  93. 93. Kollien AH, Schmidt J, Schaub GA (1998) Modes of association of Trypanosoma cruzi with the intestinal tract of the vector Triatoma infestans. Acta Trop 70: 127–141.
  94. 94. Killick Kendrick R, Molyneux DH, Ashford RW (1974) Leishmania in phlebotomid sandflies. I. Modifications of the flagellum associated with attachment to the mid gut and oesophageal valve of the sandfly. Proc R Soc B 187: 409–419.
  95. 95. Killick-Kendrick R, Wallbanks KR, Molyneux DH, Lavin DR (1988) The ultrastructure of Leishmania major in the foregut and proboscis of Phlebotomus papatasi. Parasitology Research 74: 586–590
  96. 96. Wakid MH, Bates PA (2004) Flagellar attachment of Leishmania promastigotes to plastic film in vitro. Exp Parasitol 106: 173–178
  97. 97. Brooks AS (1978) Ultrastructure of the Flagellar Attachment Site in Three Species of Trypanosomatids. Transactions of the American Microscopical Society 97: 287–296
  98. 98. Tetley L, Vickerman K (1985) Differentiation in Trypanosoma brucei: host-parasite cell junctions and their persistence during acquisition of the variable antigen coat. J Cell Sci 74: 1–19.
  99. 99. Gregory TR (2005) Erythrocyte Cell Size Database. http://www.genomesize.com/cellsize/. Erythrocyte Cell Size Database. Available: http://www.genomesize.com/cellsize/. Accessed 23 August 2012.
  100. 100. Jastrzebski M (1894) Polymorphism of flagellates Trypanosoma Gruby, 1841, occurring in blood of crucian carp (Carassius auratus gibelio Bloch), perch (Perca fluviatilis L.) and stone perch (Acerina cernua L.). Wiadomosci Parazytologiczne 30: 172–182.
  101. 101. Hanna W (1903) Memoirs: Trypanosoma in Birds in India. Quarterly Journal of Microscopical Science s2–47: 433–438.
  102. 102. Kendall AI (1906) A new species of trypanosome occurring in the mouse Mus musculus. Journal of Infectious Diseases 3: 228–231.
  103. 103. Yakimoff WL (1918) Trypanosomiasis of Camels in Russian Turkestan. Parasitology 11: 35–80
  104. 104. Hoare CA (1923) An Experimental Study of the Sheep-Trypanosome (T. melophagium Flu, 1908), and its Transmission by the Sheep-Ked (Melophagus ovinus L.). Parasitology 15: 365–424
  105. 105. Channon HA, Wright HD (1927) Observations on trypanosomiasis of rabbits, and its natural mode of transmission. The Journal of Pathology and Bacteriology 30: 253–260.
  106. 106. Nigrelli RF (1929) On the Cytology and Life-History of Trypanosoma diemyctyli and the Polynuclear Count of Infected Newts (Triturus viridescens). Transactions of the American Microscopical Society 48: 366
  107. 107. Fantham HB, Porter A, Richardson LR (1942) Some Haematozoa observed in vertebrates in eastern Canada. Parasitology 34: 199–226
  108. 108. Lehmann DL (1954) A New Species of Trypanosome from the Salamander Ambystoma gracile, with Notes on a Collection of Amphibian Blood Smears. The Journal of Parasitology 40: 656
  109. 109. Saunders DC (1959) Trypanosoma balistes n. sp. from Balistes capriscus Gmelin, the Common Triggerfish, from the Florida Keys. The Journal of Parasitology 45: 623
  110. 110. Qadri SS (1962) On three new trypanosomes from freshwater fishes. Parasitology 52: 221–228
  111. 111. Shaw JJ, Lainson R (1972) Trypanosoma vivax in Brazil. Ann Trop Med Parasitol 66: 25–33.
  112. 112. Kingston N, Morton JK (1975) Trypanosoma cervi sp. n. from Elk (Cervus canadensis) in Wyoming. The Journal of Parasitology 61: 17–23
  113. 113. Werner JK, Walewski K (1976) Amphibian Trypanosomes from the McCormick Forest, Michigan. The Journal of Parasitology 62: 20
  114. 114. Letch CA (1979) Host restriction, morphology and isoenzymes among trypanosomes of some British freshwater fishes. Parasitology 79: 107–117
  115. 115. Kingston N, Thorne ET, Thomas GM, McHolland L, Trueblood MS (1981) Further studies on trypanosomes in game animals in Wyoming II. J Wildlife Dis 17: 539–546.
  116. 116. Barta JR, Desser SS (1984) Blood parasites of amphibians from Algonquin Park, Ontario. J Wildlife Dis 20: 180–189.
  117. 117. Weinman D, White EA, Antipa GA (1984) Trypanosoma lucknowi, a New Species of Trypanosome from Macaca mulatta with Observations on Its Fine Structure. Journal of Eukaryotic Microbiology 31: 429–433
  118. 118. Woo PTK, Black GA (1984) Trypanosoma danilewskyi: Host Specificity and Host’s Effect on Morphometrics. The Journal of Parasitology 70: 788
  119. 119. John MC, Nedunchelliyan S, Venkataraman KS (1992) Biometrical observations on different strains of Trypanosoma evansi. Veterinary Parasitology 43: 143–145
  120. 120. Jones SRM, Woo PTK (1993) Biology of Trypanosoma burresoni n. sp. from the American Eel, Anguilla rostrata. The Journal of Parasitology 79: 542–547
  121. 121. Werner JK (1993) Blood Parasites of Amphibians from Sichuan Province, People’s Republic of China. The Journal of Parasitology 79: 356–363
  122. 122. Jones SRM, Woo PTK (1994) Morphology and Infectivity of Cultivated Trypanosoma ambystomae. The Journal of Parasitology 80: 521
  123. 123. Miyata A, Yong HS (1994) Four New Trypanosomes (Protozoa: Trypanosomatidae) in The Blood Of A Malaysian Frog, Rana blythi (Amphibia: Ranidae). 44: 539–550.
  124. 124. Nandi NC, Bennett GF, Nandi NC, Bennett GF (1994) Re-description of Trypanosoma corvi Stephens and Christophers,1908 Emend. Baker, 1976 and remarks on the Trypanosomes of the Avian Family Corvidae. Memórias do Instituto Oswaldo Cruz 89: 145–151
  125. 125. Bennett GF, Siikamäki P, Rätti O, Allander K, Gustafsson L, et al. (1994) Trypanosomes of some Fennoscandian birds. Memórias do Instituto Oswaldo Cruz 89: 531–537
  126. 126. Ziccardi M, Lourenço-de-Oliveira R (1998) Morphological Features of Trypanosomes from Squirrel Monkeys from the Brazilian Amazon. Memórias do Instituto Oswaldo Cruz 93: 301–301
  127. 127. Rodrigues AC, Campaner M, Takata CSA, Dell’ Porto A, Milder R, et al. (2003) Brazilian isolates of Trypanosoma (Megatrypanum) theileri: diagnosis and differentiation of isolates from cattle and water buffalo based on biological characteristics and randomly amplified DNA sequences. Veterinary Parasitology 116: 185–207
  128. 128. Karbowiak G, Wita I (2004) Trypanosoma (Herpetosoma) grosi kosewiense subsp. n., the parasite of the yellow-necked mouse Apodemus flavicollis (Melchior, 1834). Acta protozoologica 43: 173–178.
  129. 129. Karlsbakk E, Nylund A (2006) Trypanosomes infecting cod Gadus morhua L. in the North Atlantic: a resurrection of Trypanosoma pleuronectidium Robertson, 1906 and delimitation of T. murmanense Nikitin, 1927 (emend.), with a review of other trypanosomes from North Atlantic and Mediterranean teleosts. Systematic Parasitology 65: 175–203
  130. 130. Sehgal RNM, Valkiūnas G, Iezhova TA, Smith TB (2006) Blood parasites of chickens in Uganda and Cameroon with molecular descriptions of Leucocytozoon schoutedeni and Trypanosoma gallinarum. Journal of Parasitology 92: 1336–1343.
  131. 131. Yeld EM, Smit NJ (2006) A new species of Trypanosoma (Kinetoplastida: Trypanosomatidae) infecting catsharks from South Africa. Journal of the Marine Biological Association of the United Kingdom 86: 829–833
  132. 132. Sato H, Al-Adhami B, Une Y, Kamiya H (2007) Trypanosoma (Herpetosoma) kuseli sp. n. (Protozoa: Kinetoplastida) in Siberian flying squirrels (Pteromys volans). Parasitology Research 101: 453–461
  133. 133. Lainson R, Da Silva FMM, Franco CM (2008) Trypanosoma (Megatrypanum) saloboense n. sp. (Kinetoplastida: Trypanosomatidae) parasite of Monodelphis emiliae (Marsupiala: Didelphidae) from Amazonian Brazil. Parasite 15: 99–103.
  134. 134. Mo X, Zhang H, Jiang H, Tan Y, Shi W, et al.. (2008) Survey on the Prevalence and Morphology of Trypanosoma lewisi in Wild Rats of Guangxi Areas. Journal of Tropical Medicine 8: 621–622,597.
  135. 135. Austen JM, Jefferies R, Friend JA, Ryan U, Adams P, et al. (2009) Morphological and molecular characterization of Trypanosoma copemani n. sp. (Trypanosomatidae) isolated from Gilbert’s potoroo (Potorous gilbertii) and quokka (Setonix brachyurus). Parasitology 136: 783–792
  136. 136. Karbowiak G, Stanko M, Fričová J, Wita I, Hapunik J, et al. (2009) Blood parasites of the striped field mouse Apodemus agrarius; and their morphological characteristics. Biologia 64: 1219–1224
  137. 137. McInnes LM, Gillett A, Ryan UM, Austen J, Campbell RSF, et al. (2009) Trypanosoma irwini n. sp (Sarcomastigophora: Trypanosomatidae) from the koala (Phascolarctos cinereus). Parasitology 136: 875–885
  138. 138. Tamarit A, Tejedor-Junco MT, González M, Alberola J, Gutierrez C (2011) Morphological and biometrical features of Trypanosoma evansi isolates from an outbreak in mainland Spain. Veterinary Parasitology 177: 152–156
  139. 139. Valkiūnas G, Iezhova TA, Carlson JS, Sehgal RNM (2011) Two New Trypanosoma Species from African Birds, with Notes on the Taxonomy of Avian Trypanosomes. Journal of Parasitology 97: 924–930
  140. 140. Gupta D, Gupta N, Gangwar R (2012) Infectivity of Bufo melanostictus (Amphibia: Bufonidae) to Two New Species of Haematozoan Parasites from Rohilkhand, India. Proceedings of the Zoological Society 65: 22–32
  141. 141. Malassez M (1872) De la numération des globules rouges du sang chez les mammifères, les oisseaux et les poissons. Comptes Rendus de l’Académie des Sciences 75: 1528–1531.
  142. 142. Gulliver G (1875) Observations on the sizes and shapes of the red corpuscles of the blood of vertebrates, with drawings of them to a uniform scale, and extended and revised tables of measurements. Proceedings of the Zoological Society of London: 474–495.
  143. 143. Cleland JB, Johnston TH (1912) Relative dimensions of the red blood cells of vertebrates, especially of birds. Emu 11: 188–197.
  144. 144. Smith HM (1925) Cell size and metabolic activity in Amphibia. Biological Bulletin 48: 347–378.
  145. 145. Emmons WF (1927) The interrelation of number, volume, diameter and area of mammalian erythrocytes. The Journal of Physiology 64: 215–228.
  146. 146. Wintrobe MM (1934) Variations in the size and hemoglobin content of erythrocytes in the blood of various vertebrates. Folia Haematologica 51: 32–49.
  147. 147. Bartsch P, Rosenzweig W (1937) Size of red blood corpuscles and their nucleus in fifty North American birds. The Auk 54: 516–519.
  148. 148. Ponder E (1948) Hemolysis and related phenomena. Grune & Stratton. 416 p.
  149. 149. Kisch B (1949) Hemoglobin content, size and amount of erythrocytes in fishes. Experimental medicine and surgery 7: 118.
  150. 150. Kisch B (1949) Observations on the haematology of fishes and birds. Experimental medicine and surgery 7: 318.
  151. 151. Kisch B (1951) Erythrocytes in fishes. Experimental medicine and surgery 9: 125.
  152. 152. Data AI of BSC on the H of B (1952) Standard values in blood: being the first fascicle of a handbook of biological data. Saunders. 222 p.
  153. 153. Vernberg FJ (1955) Hematological studies on salamanders in relation to their ecology. Herpetologica 11: 129–133.
  154. 154. Hartman FA, Lessler MA (1963) Erythrocyte measurements in birds. The Auk 80: 467–473.
  155. 155. Hartman FA, Lessler MA (1964) Erythrocyte measurements in fishes, amphibia and reptiles. The Biological Bulletin 126: 83–88.
  156. 156. Saunders DC (1966) Elasmobranch blood cells. Copeia 1966: 348–351.
  157. 157. Szarski H, Czopek G (1966) Erythrocyte diameter in some amphibians and reptiles. Bull Acad Pol Sci Cl II Ser Sci Biol 14: 433–437.
  158. 158. Conroy DA, Rodriguez JL (1966) Erythrocyte Measurements of Some Argentine Fishes. The Progressive Fish-Culturist 28: 46–46.
  159. 159. Rouf MA (1969) Hematology of the leopard frog, Rana pipiens. Copeia: 682–687.
  160. 160. Goniakowska L (1970) The respiration of erythrocytes of some amphibias in vitro. Bulletin de l’Académie polonaise des sciences Série des sciences biologiques 18: 793.
  161. 161. Monnickendam MA, Balls M (1973) The relationship between cell sizes, respiration rates and survival of amphibian tissues in long-term organ cultures. Comparative Biochemistry and Physiology Part A: Physiology 44: 871–880.
  162. 162. Hawkey CM (1975) Comparative mammalian haematology: cellular components and blood coagulation of captive wild animals. Heinemann Medical. 332 p.
  163. 163. Mitsuru K (1981) Relationships between number, size and shape of red blood cells in amphibians. Comparative Biochemistry and Physiology Part A: Physiology 69: 771–775.
  164. 164. Lewis JH (1996) Comparative hemostasis in vertebrates. Plenum Press. 456 p.
  165. 165. Benga G, Kuchel PW, Chapman BE, Cox GC, Ghiran I, et al. (2000) Comparative cell shape and diffusional water permeability of red blood cells from Indian elephant (Elephas maximus) and Man (Homo sapiens). Comparative Haematology International 10: 1–8.
  166. 166. Gregory TR (2000) Nucleotypic effects without nuclei: genome size and erythrocyte size in mammals. Genome 43: 895–901.
  167. 167. Gentry PA (2004) Comparative aspects of blood coagulation. The Veterinary Journal 168: 238–251.
  168. 168. Collins TJ (2007) ImageJ for microscopy. BioTechniques 43: 25–30.
  169. 169. Mackinnon DL (1909) Note on two new flagellate parasites in fleas - Herpetomonas ctenophthalmi, n. sp., and Crithidia hystrichopsyllae, n. sp. Parasitology 2: 288–296.
  170. 170. Mackinnon DL (1911) On some more protozoan parasites from Trichoptera. Parasitology 4: 28–38.
  171. 171. Fantham HB (1912) Herpetomonas pediculi, nov. spec., parasitic in the alimentary tract of Pediculus vestimenti, the human body louse. Proceedings of the Royal Society of London Series B, Containing Papers of a Biological Character 84: 505–517.
  172. 172. Tyzzer EE, Walker EL (1919) A Comparative Study of Leishmania Infantum of Infantile Kala Azar and Leptomonas (Herpetomonas) Ctenocephali Parasitic in the Gut of the Dog Flea. J Med Res 40: 129–176.5.
  173. 173. Noguchi H, Tilden EB (1926) Comparative Studies of Herpetomonads and Leishmanias. J Exp Med 44: 307–325.
  174. 174. Becker ER (1927) Herpetomonas arctocorixae sp. nov. Entozoic in the Intestine of the Water-Boatman Arctocorixa interrupta Say. The Journal of Parasitology 14: 85–87
  175. 175. Wallace FG (1943) Flagellate Parasites of Mosquitoes with Special Reference to Crithidia fasciculata Léger, 1902. The Journal of Parasitology 29: 196–205
  176. 176. Wallace FG (1962) The Trypanosomatid Parasites of Horseflies with the Description of Crithidia rileyi n. sp. Journal of Eukaryotic Microbiology 9: 53–58
  177. 177. Gibbs AJ (1947) Observations on leptomonas ctenocephali (Fantham, 1912). Transactions of the Royal Society of Tropical Medicine and Hygiene 40: 495–500.
  178. 178. Gibbs AJ (1951) Leptomonas capsularis n. sp. and other flagellates parasitic in Cletus ochraceus (Hemiptera). Parasitology 48: 128–133.
  179. 179. Gibbs AJ (1959) Leptomonas sp., parasitic in the fly Leia sp. (fam. Mycetophilidae), with special reference to the metacyclic forms. Parasitology 49: 411–415.
  180. 180. Wallace FG, Clark TB, Dyer MI, Collins T (1960) Two New Species of Flagellates Cultivated from Insects of the Genus Gerris. Journal of Eukaryotic Microbiology 7: 390–392
  181. 181. Wallace FG, Todd SR, Rogers W (1965) Flagellate Parasites of Water Striders with a Description of Leptomonas costoris, n. sp. Journal of Eukaryotic Microbiology 12: 390–393.
  182. 182. Hanson WL, McGhee RB (1961) The Biology and Morphology of Crithidia acanthocephali n. sp., Leptomonas leptoglossi n. sp., and Blastocrithidia euschisti n. sp. Journal of Eukaryotic Microbiology 8: 200–204
  183. 183. Gillies C, Hanson ED (1963) A New Species of Leptomonas Parasitizing the Macronucleus of Paramecium trichium. Journal of Eukaryotic Microbiology 10: 467–473.
  184. 184. Langridge DF, McGHEE RB (1967) Crithidia mellificae n. sp. an Acidophilic Trypanosomatid of the Honey Bee Apis mellifera. Journal of Eukaryotic Microbiology 14: 485–487
  185. 185. McGhee RB, Hanson WL, Schmittner SM (1969) Isolation, Cloning and Determination of Biologic Characteristics of Five New Species of Crithidia. Journal of Eukaryotic Microbiology 16: 514–520
  186. 186. Smirnoff WA, Lipa JJ (1970) Herpetomonas swainei sp. n., a new flagellate parasite of Neodiprion swainei (Hymenoptera: Tenthredinidae). Journal of Invertebrate Pathology 16: 187–195.
  187. 187. Rogers WE, Wallace FG (1971) Two new subspecies of Herpetomonas muscarum (Leidy, 1856) Kent, 1880. Journal of Eukaryotic Microbiology 18: 645–649.
  188. 188. Daggett PM, Dollahon N, Janovy Jr J (1972) Herpetomonas megaseliae sp. n.(Protozoa: Trypanosomatidae) from Megaselia scalaris (Loew, 1866) Schmitz, 1929 (Diptera: Phoridae). The Journal of Parasitology: 946–949.
  189. 189. Roitman I, Mundim MH, Azevedo HPD, Kitajima EW (1977) Growth of Crithidia at High Temperature: Crithidia hutneri sp. n. and Crithidia luciliae thermophila s. sp. n. Journal of Eukaryotic Microbiology 24: 553–556
  190. 190. Yoshida N, Freymüller E, Wallace FG (1978) Herpetomonas mariadeanei sp. n. (Protozoa, Trypanosomatidae) from Muscina stabulans (Falléen, 1816) (Diptera, Muscidae). Journal of Eukaryotic Microbiology 25: 421–425
  191. 191. De Sá MFG, De Sá CM, Veronese MA, Filho SA, Gander ES (1980) Morphologic and Biochemical Characterization of Crithidia Brasiliensis Sp. N. Journal of Eukaryotic Microbiology 27: 253–257
  192. 192. Manaia ADAC, Souza M, Do Carmo Moreira D, Lustosa ÉDES, Roitman I (1981) Leptomonas lactosovorans n. sp., a Lactose-Utilizing Trypanosomatid: Description and Nutritional Requirements1. Journal of Eukaryotic Microbiology 28: 124–126.
  193. 193. Malone LA, Wigley PJ, Dhana SD (1985) A flagellate protozoan from Mythimna separata (Lepidoptera: Noctuidae). New Zealand Journal of Zoology 12: 155–157
  194. 194. Beard CB, Butler JF, Greiner EC (1989) In vitro Growth Characterization and Host-Parasite Relationship of Leptomonas pulexsimulantis n. sp., a Trypanosomatid Flagellate of the Flea Pulex simulans. The Journal of Parasitology 75: 658–668
  195. 195. Jankevicius JV, Jankevicius SI, Campaner M, Conchon I, Maeda LA, et al. (1989) Life Cycle and Culturing of Phytomonas serpens (Gibbs), a Trypanosomatid Parasite of Tomatoes. Journal of Eukaryotic Microbiology 36: 265–271
  196. 196. Motta MCM, Cava AMS, Silva PMF, Fiorini JE, Soares MJ, et al. (1991) Morphological and biochemical characterization of the trypanosomatids Crithidia desouzai and Herpetomonas anglusteri. Canadian Journal of Zoology 69: 571–577
  197. 197. Podlipaev SA, Sturm NR, Fiala I, Fernandes O, Westenberger SJ, et al. (2004) Diversity of Insect Trypanosomatids Assessed from the Spliced Leader RNA and 5s rRNA Genes and Intergenic Regions1. Journal of Eukaryotic Microbiology 51: 283–290
  198. 198. Yurchenko VY, Lukeš J, Jirku M, Zeledón R, Maslov DA (2006) Leptomonas costaricensis sp. n. (Kinetoplastea: Trypanosomatidae), a member of the novel phylogenetic group of insect trypanosomatids closely related to the genus Leishmania. Parasitology 133: 537–546
  199. 199. Schmid-Hempel R, Tognazzo M (2010) Molecular divergence defines two distinct lineages of Crithidia bombi (Trypanosomatidae), parasites of bumblebees. J Eukaryot Microbiol 57: 337–345
  200. 200. Zídková L, Cepicka I, Votýpka J, Svobodová M (2010) Herpetomonas trimorpha sp. nov. (Trypanosomatidae, Kinetoplastida), a parasite of the biting midge Culicoides truncorum (Ceratopogonidae, Diptera). Int J Syst Evol Microbiol 60: 2236–2246
  201. 201. Borghesan TC, Ferreira RC, Takata CSA, Campaner M, Borda CC, et al.. (2012) Molecular Phylogenetic Redefinition of Herpetomonas (Kinetoplastea, Trypanosomatidae), a Genus of Insect Parasites Associated with Flies. Protist. Available: http://www.sciencedirect.com/science/article/pii/S1434461012000740. Accessed 6 September 2012.
  202. 202. Lloyd L, Johnson WB (1924) The Trypanosome Infections of Tsetse-flies in Northern Nigeria and a new Method of Estimation. Bulletin of Entomological Research 14: 265–288
  203. 203. Wenyon CM (1926) Protozoology: A Manual for Medical Men, Veterinarians and Zoologists. Baillière, Tindall and Cox. 840 p.
  204. 204. Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22: 4673–4680.
  205. 205. Sievers F, Wilm A, Dineen D, Gibson TJ, Karplus K, et al. (2011) Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol Syst Biol 7: 539
  206. 206. Lassmann T, Sonnhammer ELL (2005) Kalign–an accurate and fast multiple sequence alignment algorithm. BMC Bioinformatics 6: 298
  207. 207. Katoh K, Kuma K, Miyata T, Toh H (2005) Improvement in the accuracy of multiple sequence alignment program MAFFT. Genome Inform 16: 22–33.
  208. 208. Edgar RC (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res 32: 1792–1797
  209. 209. Notredame C, Higgins DG, Heringa J (2000) T-Coffee: A novel method for fast and accurate multiple sequence alignment. J Mol Biol 302: 205–217
  210. 210. Collingridge PW, Kelly S (2012) MergeAlign: improving multiple sequence alignment performance by dynamic reconstruction of consensus multiple sequence alignments. BMC Bioinformatics 13: 117
  211. 211. Saitou N, Nei M (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4: 406–425.
  212. 212. Lukeš J, Jirků M, Doležel D, Kral’ová I, Hollar L, et al. (1997) Analysis of ribosomal RNA genes suggests that trypanosomes are monophyletic. Journal of molecular evolution 44: 521–527.
  213. 213. Bates PA (2007) Transmission of Leishmania metacyclic promastigotes by phlebotomine sand flies. Int J Parasitol 37: 1097–1106
  214. 214. Kollien AH, Schaub GA (2003) The development of Blastocrithidia triatomae (Trypanosomatidae) in the reduviid bug Triatoma infestans (Insecta): influence of feeding. Parasitol Res 89: 430–436
  215. 215. Laird M (1959) Blastocrithidia n.g. (Mastigophora: Protomonadina) for crithidia (in part) With a Subarctic Record for B. gerridis (Patton). Canadian Journal of Zoology 37: 749–752
  216. 216. Patton WS (1908) The life-cycle of a species of Crithidia parasitic in the intestinal tract of Gerris fossarum Fabr. Archiv fur Protistenkunde 12: 131–146.
  217. 217. Drbohlav JJ (1926) The Cultivation of Herpetomonas muscarum (Leidy 1856) Kent 1881 from Lucilia sericata. The Journal of Parasitology 12: 183–190
  218. 218. Wilson VCLC, Southgate BA (1979) Lizard Leishmania. Biology of the kinetoplastida. London: Academic Press, Vol. 2: 241–268.
  219. 219. Zhang LM, Leng YJ (1997) Eighty-year research of phlebotomine sandflies (Diptera: Psychodidae) in China (1915–1995). II. Phlebotomine vectors of leishmaniasis in China. Parasite 4: 299–306.
  220. 220. Elwasila M (1988) Leishmania tarentolae Wenyon, 1921 from the gecko Tarentola annularis in the Sudan. Parasitology Research 74: 591–592
  221. 221. Raymond F, Boisvert S, Roy G, Ritt J-F, Légaré D, et al.. (2011) Genome sequencing of the lizard parasite Leishmania tarentolae reveals loss of genes associated to the intracellular stage of human pathogenic species. Nucleic Acids Research. Available: http://www.ncbi.nlm.nih.gov/pubmed/21998295. Accessed 18 October 2011.
  222. 222. Vaughan S, Kohl L, Ngai I, Wheeler RJ, Gull K (2008) A Repetitive Protein Essential for the Flagellum Attachment Zone Filament Structure and Function in Trypanosoma brucei. Protist 159: 127–136
  223. 223. Woods K, Nic a’Bhaird N, Dooley C, Perez-Morga D, Nolan DP (2013) Identification and characterization of a stage specific membrane protein involved in flagellar attachment in Trypanosoma brucei. PLoS ONE 8: e52846
  224. 224. Oberholzer M, Langousis G, Nguyen HT, Saada EA, Shimogawa MM, et al. (2011) Independent analysis of the flagellum surface and matrix proteomes provides insight into flagellum signaling in mammalian-infectious Trypanosoma brucei. Mol Cell Proteomics 10: M111.010538
  225. 225. Galetović A, Souza RT, Santos MRM, Cordero EM, Bastos IMD, et al. (2011) The repetitive cytoskeletal protein H49 of Trypanosoma cruzi is a calpain-like protein located at the flagellum attachment zone. PLoS ONE 6: e27634
  226. 226. Hyams JS (1982) The Euglena paraflagellar rod: structure, relationship to other flagellar components and preliminary biochemical characterization. J Cell Sci 55: 199–210.
  227. 227. Woodward R, Carden MJ, Gull K (1994) Molecular characterisation of a novel, repetitive protein of the paraflagellar rod in Trypanosoma brucei. Mol Biochem Parasitol 67: 31–39.
  228. 228. Pullen TJ, Ginger ML, Gaskell SJ, Gull K (2004) Protein targeting of an unusual, evolutionarily conserved adenylate kinase to a eukaryotic flagellum. Mol Biol Cell 15: 3257–3265
  229. 229. Portman N, Lacomble S, Thomas B, McKean PG, Gull K (2009) Combining RNA interference mutants and comparative proteomics to identify protein components and dependences in a eukaryotic flagellum. J Biol Chem 284: 5610–5619
  230. 230. Hertz-Fowler C, Ersfeld K, Gull K (2001) CAP5.5, a life-cycle-regulated, cytoskeleton-associated protein is a member of a novel family of calpain-related proteins in Trypanosoma brucei. Mol Biochem Parasitol 116: 25–34
  231. 231. Olego-Fernandez S, Vaughan S, Shaw MK, Gull K, Ginger ML (2009) Cell morphogenesis of Trypanosoma brucei requires the paralogous, differentially expressed calpain-related proteins CAP5.5 and CAP5.5V. Protist 160: 576–590
  232. 232. Rindisbacher L, Hemphill A, Seebeck T (1993) A repetitive protein from Trypanosoma brucei which caps the microtubules at the posterior end of the cytoskeleton. Molecular and Biochemical Parasitology 58: 83–96
  233. 233. Baines A, Gull K (2008) WCB is a C2 domain protein defining the plasma membrane - sub-pellicular microtubule corset of kinetoplastid parasites. Protist 159: 115–125
  234. 234. Bonhivers M, Nowacki S, Landrein N, Robinson DR (2008) Biogenesis of the Trypanosome Endo-Exocytotic Organelle Is Cytoskeleton Mediated. PLoS Biol 6: e105
  235. 235. Morriswood B, He CY, Sealey-Cardona M, Yelinek J, Pypaert M, et al. (2009) The bilobe structure of Trypanosoma brucei contains a MORN-repeat protein. Mol Biochem Parasitol 167: 95–103
  236. 236. Zhou Q, Gheiratmand L, Chen Y, Lim TK, Zhang J, et al. (2010) A comparative proteomic analysis reveals a new bi-lobe protein required for bi-lobe duplication and cell division in Trypanosoma brucei. PLoS ONE 5: e9660
  237. 237. He CY, Pypaert M, Warren G (2005) Golgi duplication in Trypanosoma brucei requires Centrin2. Science 310: 1196–1198
  238. 238. Sun SY, Wang C, Yuan YA, He CY (2013) An intracellular membrane junction consisting of flagellum adhesion glycoproteins links flagellum biogenesis to cell morphogenesis in Trypanosoma brucei. J Cell Sci 126: 520–531
  239. 239. Müller N, Hemphill A, Imboden M, Duvallet G, Dwinger RH, et al. (1992) Identification and characterization of two repetitive non-variable antigens from African trypanosomes which are recognized early during infection. Parasitology 104: 111–120
  240. 240. Sherwin T, Gull K (1989) The cell division cycle of Trypanosoma brucei brucei: timing of event markers and cytoskeletal modulations. Philos Trans R Soc Lond, B, Biol Sci 323: 573–588.