Flagellar motility in eukaryotic human parasites

G Model

ARTICLE IN PRESS

YSCDB-1865; No. of Pages 15

Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

Contents lists available at ScienceDirect

Seminars in Cell & Developmental Biology journal homepage: www.elsevier.com/locate/semcdb

Review

Flagellar motility in eukaryotic human parasites Timothy Krüger, Markus Engstler ∗ Department of Cell and Developmental Biology, Biocentre, University of Wuerzburg, Am Hubland, 97074 Wuerzburg, Germany

a r t i c l e

i n f o

Article history: Received 2 September 2015 Received in revised form 26 October 2015 Accepted 26 October 2015 Available online xxx Keywords: Microbial motility Flagellum Unicellular parasite Spatiotemporal resolution Trypanosoma Giardia Trichomonas Leishmania Plasmodium

a b s t r a c t A huge variety of protists rely on one or more motile flagella to either move themselves or move fluids and substances around them. Many of these flagellates have evolved a symbiotic or parasitic lifestyle. Several of the parasites have adapted to human hosts, and include agents of prevalent and serious diseases. These unicellular parasites have become specialised in colonising a wide range of biological niches within humans. They usually have diverse transmission cycles, and frequently manifest a variety of distinct morphological stages. The motility of the single or multiple flagella plays important but understudied roles in parasite transmission, host invasion, dispersal, survival, proliferation and pathology. In this review we provide an overview of the important human pathogens that possess a motile flagellum for at least part of their life cycle. We highlight recently published studies that aim to elucidate motility mechanisms, and their relevance for human disease. We then bring the physics of swimming at the microscale into context, emphasising the importance of interdisciplinary approaches for a full understanding of flagellate motility – especially in light of the parasites’ microenvironments and population dynamics. Finally, we summarise some important technological aspects, describing challenges for the field and possibilities for motility analyses in the future. © 2015 Elsevier Ltd. All rights reserved.

Contents 1. 2.

3.

4. 5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Mechanisms of motility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 2.1. The axoneme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 2.2. Flagellar and ciliary beats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Flagellar parasites of humans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.1. Metamonads . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.1.1. Trichomonads – Trichomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.1.2. Diplomonads – Giardia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.2. Kinetoplastids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.2.1. Leishmania . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.2.2. Trypanosoma cruzi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.2.3. Trypanosoma brucei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 3.3. Apicomplexa – Plasmodium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Physics of flagellated microswimmers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Technical perspective and challenges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.1. Spatiotemporal resolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.2. The third dimension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.3. Tissue models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.4. In vivo analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.5. Molecular genetic tools . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00

∗ Corresponding author. E-mail address: [email protected] (M. Engstler). http://dx.doi.org/10.1016/j.semcdb.2015.10.034 1084-9521/© 2015 Elsevier Ltd. All rights reserved.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

2

1. Introduction Flagellar motility is an ancestral eukaryotic concept. It is widely accepted that the last eukaryotic common ancestor featured a motile, microtubule-scaffolded organelle powered by dynein motor proteins [1]. Consequently, flagellates are widespread across eukaryotic supergroups [2,3]. Flagellates, possessing one or several of these snakelike appendages, are free-swimming or sessile protists, and can be either free-living or symbiotic. They are a major constituent of the global planktonic microbiome, whose huge diversity is currently being charted [4]. Flagellates have also been recognised as important parasites of animals and humans since the late nineteenth century [5]. Flagellate parasites of humans belong either to the group Metamonada, which are amitochondriate, tetrakont zooflagellates [6], including Giardia and Trichomonas, or to the kinetoplastids (Fig. 1). Kinetoplastids are characterised by their eponymous mitochondrial DNA structure. They may be the

earliest-branching group of parasitic protists, and have a very broad distribution of animal and plant hosts [7]. Kinetoplastid parasites of humans belong to the genera Trypanosoma and Leishmania. In addition, the almost entirely parasitic Apicomplexa rely on flagellate male microgametes during sexual reproduction. The group contains several human- and livestock-infecting genera, amongst them Plasmodium and Toxoplasma, which form cell-invasive asexual sporozoites during their life cycle [8]. Finally, another human parasite has to be mentioned, Naegleria fowleri, a free-living amoeboflagellate that feeds and divides in an amoeboid life-cycle form. In this trophozoite stage, Naegleria infects the central nervous system, albeit rarely but with deadly outcome [9,10]. Naegleria trophozoites have the fascinating ability to change from the amoeboid form into a flagellate by developing two flagella de novo, together with a complete microtubule cytoskeleton and basal bodies [11]. Free-living flagellates display a tremendous morphological variety (e.g. [12]), reflecting their vast range of habitats, lifestyles, and

Fig. 1. Flagellate morphotypes of human eukaryotic parasites. Left: The evolutionary position of the parasites (coloured boxes) is shown, based on the classification of the five boxed super-groups according to [3] and comparing several other flagellate species and groups. Typical views of flagellate forms are shown in comparison (for dimensions see Table 1). CEA = common eukaryotic ancestor, SAR = Stramenopiles, Alveolates and Rhizaria. Yellow shading highlights the parasitic flagellates that are discussed in this review, while the classical model systems Chlamydomonas and sperm are shown in the lower box.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

evolutionary histories. In many cases, the purpose and advantage of flagellar motility is intuitively clear. Free-swimming protists can orientate towards the light or prey to acquire resources, towards mating partners for sexual reproduction, or move away from other organisms for safety or dispersal. Stationary freeliving protozoa use their flagella to produce hydrodynamic flows in order to improve resource uptake [13]. In contrast, in most cases of parasitic flagellates it is not entirely clear what constraints shape their motile behaviour. The parasites must adapt to distinct, hostile, and largely confined microenvironments inside their hosts. Besides specific immunological and biochemical challenges therein, they also face considerable physical challenges and barriers. They have evolved distinct developmental adaptations of metabolism and morphology in order to optimise their survival in various host environments, and facilitate their successful transmission between different host animals [14]. Integrative approaches, on the single cell level, are being undertaken in order to unravel the complex host–parasite interactions that ensue [15,16]. Consequently, the physics of protist motility has come into interdisciplinary focus, with the goal of generating a complete and quantitative picture of the environmental parameters relevant for specific cells’ behaviour and survival [17]. While the general biology of infective cycles is relatively well understood, especially for human parasites, details of flagellum-associated virulence remain enigmatic and are technically demanding to investigate. In this review, we summarise the lifestyles of the flagellate parasites relevant to humans, with special emphasis on their motility as a virulence factor. We briefly consider current physical understanding of microswimming, and assess extant technologies for the integrative analysis of motile parasitic behaviour and its relevance for interactions with the host. First, however, the mechanisms of flagellar motility will be outlined.

2. Mechanisms of motility 2.1. The axoneme The motor of all eukaryotic flagellar motility is the axoneme, usually in its canonical “92” form. This consists of nine microtubule doublets, cylindrically arranged and connected equidistantly to lie parallel to the long axis of the flagellum. Two single parallel microtubules are fastened longitudinally in the centre axis of the cylinder by spokes and associated protein assemblies, in all consisting of more than 250 proteins. Amongst these are the force-producing dynein motor proteins, arranged in rows of outer arms and with inner dynein arms between adjacent doublets. The tethered motor proteins are alternately activated on opposite sides of the axoneme and thus bend the elastically interconnected structure to one side or the other; the beat plane is defined by the central microtubule pair. The resulting oscillatory bending is called the flagellar beat, which is propagated along the axoneme to produce a propulsive flagellar wave. Although the mechanical aspects of the axoneme are thought to be fairly well understood, the mechanisms controlling beat cycles and coordinating wave progression are still unclear [18–21]. Flagellar proteomes have been determined in various model organisms and are comparable with regard to the conserved core axonemal structures and their function in motility. Altogether, around 650 flagellar proteins have been identified and are undergoing analysis of functions in several additional roles including sensing, signalling and intraflagellar transport, as well as attachment [22]. The biflagellate green alga Chlamydomonas reinhardtii is an important reference [23], which can be compared to human sperm [24] and the parasite Trypanosoma brucei [25–27].

3

Many flagellate axonemes are accompanied by accessory structures, which have typically been described decades ago but whose precise function remains ill-defined. Examples are the paraflagellar rod (PFR) in trypanosomatids (see Section 3.2) and microtubular structures and fibres in multiflagellate parasites. The multiflagellate parasites possess free and attached flagella on the same cell body, which complicates analyses of their relative contributions to the organism’s motility (see Section 3.1). 2.2. Flagellar and ciliary beats Axonemal oscillations are the basis for the motility of flagella and cilia. There is no structural difference between the axonemes of motile cilia and flagella. Motile cilia are shorter than flagella and usually found in larger numbers, beating in coordinated, metachronal patterns, as in ciliates and ciliated epithelia of the animal body [28]. In contrast to flagella, which often produce continuous, symmetrical and sinusoidal waves, as in sperm, motile cilia produce alternating fast, straight power strokes and slower, bending recovery strokes [17]. Some flagella can switch to perform alternating similar power and recovery strokes, which are therefore called ciliary beats, as exemplified in the green algae Chlamydomonas. This model flagellate has two free anterior flagella which beat symmetrically, performing a propulsive power stroke followed by a recovery stroke which results in a “breaststroke” swimming mechanism [29]. Chlamydomonas axonemes can switch to a flagellar waveform which results in backward movement [30]. Multiflagellated parasites can use both flagellar and ciliary beats with different types of flagella for specific functions (see Section 3.1.2). The monoflagellate trypanosomatids regularly switch between different flagellar waveforms with their single axoneme. Whereas forward movement is caused by a symmetrical flagellar wave, running from the distal tip to the proximal base, spontaneous reversal to an asymmetric base-to-tip wave can occur (see Section 3.2.3). In morphotypes with a relatively short flagellum, the reverse flagellar movement resembles, and is called, a ciliary beat [31]. Conversely, in longer flagella the reverse beat actually produces a smooth wave [32]. 3. Flagellar parasites of humans We will now consider the different flagellate parasites of humans, and the contributions their flagella make to their life cycles. 3.1. Metamonads The metamonads are characterised by having four basal bodies, which give rise to a typical pattern of three anterior and one posterior flagella [6]. They include widespread symbionts of animals, and common human parasites. 3.1.1. Trichomonads – Trichomonas Trichomonads are found in the urogenital tract, oral cavity, respiratory and digestive tract of infected humans [33]. Trichomonas vaginalis is the best characterised species, as it is responsible for the sexually transmitted infection with the highest global prevalence [34]. Four anterior flagella originate from the basal bodies of Trichomonas, which are combined into the axostylepelta complex, consisting of cross-linked microtubular structures and characteristic for the metamonad class Parabasalia. In addition to the four freely beating anterior flagella, trichomonads have one flagellum attached at several points along the cell body. This forms an undulating membrane consisting of and connected to various poorly defined fibrous structures in a species-specific manner (Fig. 2) [35,36].

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15 4

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

Fig. 2. The basic morphological features of Trichomonas vaginalis.

Fig. 3. Morphology of Giardia lamblia.

One outstanding cell biological feature of Trichomonas is the transition from the flagellar free-swimming morphotype (the trophozoite) to an amoeboid cell stage, which occurs just minutes after contact with vaginal epithelial cells. This transition is complete in a few minutes and is thus the fastest known, quicker than the common textbook example of Naegleria gruberi [11]. Although this transformation has been analysed on the molecular level [37], the actual motility of the trophozoite has largely been neglected. The flagellar apparatus is not degraded in the amoeboid stage and the cell can apparently rapidly detach and change location. Motility behaviour analyses thus seem timely, given the complex interactions of the parasite with the vaginal mucosa and microbiota, as well as its life cycle in the highly variable fluid environments of urogenital tracts [38]. Putative collective behaviour in settings of various viscosity [39] could render Trichomonas an especially interesting model microswimmer [17]. To this end, Tritrichomonas foetus has recently been analysed in a high spatiotemporal resolution microscopic study, and is being promoted as a model for multi-flagellated propulsion [40]. T. foetus is a pathogen of cattle and cats, morphologically equivalent to T. vaginalis except for having one anterior flagellum less. The cytoskeleton stays rigid during swimming, allowing the analysis of the free flagella, which produce ciliary beats. The role of the undulating attached flagellum remains speculative [41,42]. The swimming patterns of single cells have been elucidated, and the forces that they produce on the level of single flagellar beats measured [40]. Quantification of the resulting hydrodynamic impact helps to explain the motility patterns of these single-celled organisms and could act as a paradigm for the analysis of chemosensory reactions. A conclusion of the study was that multiple flagella do not necessarily produce higher thrust, compared to a single sperm flagellum, for example, indicating functions in orientation or sensing. Such studies can be extended to include analyses in varying environments, in order to address further biologically relevant issues, like morphological modulation and flagellar functions in attachment.

movement respectively) allow Giardia trophozoites to quickly orientate and position themselves in the small intestine. There, they skim close to the epithelial surface, in order to find a site for attachment [44]. Giardia produces four pairs of flagella using a duplicated metamonadal body plan (Fig. 3). The axoneme pairs traverse the cytoplasm for specific distances before they become free appendages, symmetrically exiting the cell body at anteriolateral, posterolateral, ventral and caudal positions. The internal axonemal parts are associated with unique cytoplasmic structures of unknown functions [45]. Together with associated microtubule structures, the caudal flagella produce significant propulsive forces by flexing the dorsal tail of the cell, thus producing wave-like motion [45–47]. High spatiotemporal resolution microscopic analyses have allowed identification of the mechanisms of force generation in individual flagella. The lateral flagella were shown to produce ciliary power strokes similar to those of the biflagellate alga Chlamydomonas and are used for turning, whereas the ventral flagella generate typical flagellar sine wave forms for forward propulsion. The ventral flagella have been proposed to assist in near-surface manoeuvring and fluid displacement [47]. Interestingly, the morphology of the cell body influences the flagellar dynamics in a way that cooperatively generates the ideal hydrodynamic forces for evacuating fluid beneath the cell body and pushing the cell towards the intestine surface. Subsequent work has modelled the physical parameters of flagellar motility in order to precisely superimpose observed behaviour on physical data, not least in view of the potential development of microrobotic swimmers [48]. Artificial microswimmers are being tested with various propulsion mechanisms, in the hope of producing remotecontrolled machines for future biomedical applications that can efficiently navigate the viscous fluid surroundings of the body. The design of swimming methods has been inspired mainly by bacterial microorganisms [49] and could conceivably be extended and optimised with a better understanding of the propulsion mechanisms of eukaryotic flagellate parasites.

3.1.2. Diplomonads – Giardia Another very common metamonad human parasite is Giardia lamblia (syn. intestinalis, syn. duodenalis), the causative agent of one of the most prevalent gastrointestinal diseases in the world. It has two life cycle stages – a dormant cyst that is orofecally transmitted, and swimming trophozoites that are released from a cyst in the small intestine. These trophozoites attach to epithelial cells, multiply, and inflict pathogenic damage to the epithelial barrier in several ways [43]. The octoflagellate trophozoites attach to the intestinal microvilli via a ventral suction disk. The disk forms a continuous seal with the epithelial cells, preventing the invaders from being moved by peristalsis. The parasites’ flagellar motility is a key factor in the colonisation of the small intestine, as anterior and ventral flagella (responsible for rotational and forward

3.2. Kinetoplastids The kinetoplastids are divided into several clades of biflagellate bodonids and the uniflagellate Trypanosomatida. The majority of bodonids are free-swimming in plankton but have developed parasitic species several times, albeit ones almost entirely confined to aquatic hosts. They are important as evolutionary relatives of trypanosomatids, but otherwise relatively unstudied compared to their obligatory parasitic cousins [7]. The trypanosomatids represent the most diverse and adaptable group of flagellated parasites with an extremely broad host range. They include several important human and animal parasites, always being transmitted by insect vectors. This dixenous lifestyle

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

5

Fig. 4. Promastigote morphotype of Leishmania spp. The anterior (A) and posterior (P) cell poles are indicated.

goes hand in hand with a high degree of developmental plasticity. During their life cycle, the parasites undergo a species-specific succession of molecular and morphological developmental changes in order to adapt to diverse host environments. Trypanosomatid cells can usually be classified into eight morphotypes, characterised by the relative positions of the kinetoplast and the flagellum, as well as the cell shape and degree of flagellar attachment [50,51]. Most genera exhibit only a certain subset of these morphotypes, but one should keep in mind that cell forms are likely to be rather continuous and that the taxonomic value of static morphotypes is limited [7]. This could mean that the morphological variability of certain life cycle stages, especially as a function of the microenvironment, has been underestimated so far.

3.2.1. Leishmania The genus Leishmania includes at least 20 species of human pathogens, which are transmitted by several sand fly species (Phlebotomus, Lutzomyia) [52]. There are two main morphotypes, the promastigote and amastigote, which are found in the fly and human hosts, respectively. The motile parasite is the promastigote form, with a free anterior flagellum that pulls the elongated cell body along by producing planar waves (Fig. 4) [31]. The promastigote is produced in the fly directly after uptake of an amastigote with a bloodmeal. A motile flagellum is assembled by intraflagellar transport [53], emanating from the flagellar pocket at the anterior end of the cell body (Fig. 4), a characteristic membrane invagination in trypanosomatids. The motility of the promastigote plays an important role for passage from the midgut of the fly to the anterior parts of the gut [54]. This ensures that the cells are correctly positioned to be transmitted again by the insect’s bite. This translocation could be directed by chemotaxis [55]. The transmittable form of the promastigote is called a metacyclic, which is characterised by a relatively long flagellum and high motility, but more detailed information on the relevance of its motile behaviour is lacking. Besides motility, the Leishmania flagellum is especially interesting with regard to several attachment mechanisms in the sand fly gut [52]. Once inside a human host, metacyclics invade phagocytotic cells, i.e. neutrophils and macrophages, where they differentiate into the amastigote form. This process has been analysed in a high-resolution live imaging study of L. donovani, and was found to be subdivided into four phases [56]. First, the flagellar tip triggered phagocytosis by inducing pseudopod formation in the macrophage after interaction. The locomotive force produced by the flagellum towards the macrophage was shown to be essential for effective phagocytosis. The uptake of parasites was exclusively orchestrated by the flagellar tip and was significantly reduced in immobilised parasites. Second, persistent flagellar motility continued during uptake. This reorientated the flagellar tip towards the macrophage plasma membrane and the posterior end of the cell towards the host cell nucleus. Third, the parasite maintained its

intracellular position by continuing flagellar oscillations. These oscillations produced an outward directing force that equalised an inward force from the host cell. This inward force is presumably applied via the cytoskeleton to the nascent parasitophorous vacuole in which the parasite will reside. During this struggle the flagellum damaged the plasma membrane, which was repaired by concurrent lysosome exocytosis. After this, flagellar motility ceased and the cell differentiated into the immotile amastigote form. The amastigotes retain a short flagellum that remains exposed to the lumen of the parasitophorous vacuole [57–59]. This example shows how versatile the force produced by flagellar oscillations can be utilised, creating complex environment-dependent behaviour far beyond simple swimming in fluids. 3.2.2. Trypanosoma cruzi Like Leishmania spp., T. cruzi is an obligate intracellular parasite causing Chagas disease, predominantly in Latin America. The host–parasite relationship has been extensively studied on a molecular and genetic level [60,61]. The insect hosts of T. cruzi are hematophagous bugs, which take up the parasite by feeding on an infected human host. In the insect’s gastrointestinal tract, T. cruzi replicates in the epimastigote form. Epimastigotes have a flagellum that is attached to the cell body and originates at the flagellar pocket located anterior to the nucleus (Fig. 5). The anterior free part of the flagellum beats similar to that of Leishmania promastigotes, producing a symmetric tip-to-base flagellar wave, which propels the cell forward. The parasites also show beat reversals characteristic of trypanosomatids [31], producing asymmetric base-to-tip waves which interrupt forward swimming and rotate the cell [62]. In the insect gut, the epimastigotes eventually differentiate into the human-infective metacyclic trypomastigote form. Trypomastigotes are similar to epimastigotes but have a flagellar pocket positioned on the posterior side of the nucleus, resulting in a longer length of flagellum attachment. These metacyclics are deposited by the excretions of the bug in a feeding wound. The metacyclics can invade various host cell types, culminating in their uptake in lysosomal vacuoles. They subsequently escape into the

Fig. 5. Epimastigote form of Trypanosoma cruzi. The anterior (A) and posterior (P) cell poles are indicated.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15 6

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

cytoplasm where they differentiate into a replicative amastigote stage [63]. The life cycle of T. cruzi includes an additional morphotype, the bloodstream form (also of the trypomastigote type), which develops from the amastigotes in response to an unknown signal. Unlike the metacyclics, the bloodstream forms can invade non-phagocytic cells as well as macrophages. They can thus reinforce infection, as they are spread through the bloodstream and lymph of the host [63]. Flagellar motility probably plays important roles in the dispersal of the bloodstream form trypanosomes, as well as in the evasion of the immune system. Hence, motion ensures the survival of the extracellular parasites. Despite this, there are only sparse reports on challenges to the trypanosomes’ motility [64], and to date there have been no high-resolution, integrative motility studies on T. cruzi. Most research has concentrated on the invasion of host cells, and although the intracellular role of motility has tentatively been discussed for this process [65] it has not yet been rigorously tested. There is increasing interest in studying the global population variability of T. cruzi, as several discrete strains have been genetically characterised and their relevance for infectivity, virulence, disease progression and tissue tropism has been documented [66]. The nature of the preferences for various host organs, where clonal populations linger for years during chronic infections before spreading to the bloodstream again is being debated. Likewise, the effects of alternative infection routes, e.g. oral or via the gastric mucosa, are being explored [67–69]. 3.2.3. Trypanosoma brucei Most work on parasitic flagellar motility has been conducted in the African trypanosome species. These zoonotic pathogens are mostly transmitted to their diverse hosts by injection in the saliva of infected tsetse flies during their bloodmeal. In contrast to the other trypanosomatids, the African trypanosomes are strictly extracellular parasites, and retain flagellated motile morphotypes throughout their entire life cycle. Subspecies of the generally animal-infective Trypanosoma brucei – T. b. rhodesiense and T. b. gambiense – are responsible for causing disease in humans (African sleeping sickness). Their ability to infect humans is due to the development of resistance mechanisms against trypanosome lytic factors present in human serum [70,71]. Because of their otherwise identical character and behaviour, T. b. brucei has been extensively cultured and studied as the model African trypanosome. Indeed, T. b. brucei has been and continues to be an exceptional model organism, successfully allowing the discovery of several aspects of fundamental eukaryotic biology [72]. Human-infective T. brucei metacyclic trypomastigotes develop in the fly salivary glands [73]. They are formed from an asymmetric cell division of epimastigotes [74]. The metacyclics are nonproliferative, and convert into bloodstream form (BSF) trypanosomes shortly after injection into the skin. BSFs can multiply and transfer to either the bloodstream or the lymphatic system (and between these destinations) after varying periods of time [75]. There is virtually no data on the motility of metacyclics in the skin after infection and it is not clear what role this environment initially plays for BSF cell development. Chancres, indicating acute inflammatory reactions to the injection and reported as typical sites of trypanosome proliferation [76,77], are apparently only rarely seen in human infections [78]. In any case, the trypanosomes must be able to rapidly convert from the short, cell cycle-arrested metacyclic to the long slender BSF morphotype, which is capable of surviving and proliferating in the lymphatic system and the bloodstream. Flagellar motility has been suggested to be crucial for the proliferation of BSF trypanosomes, as the ablation of flagellar proteins causes cytokinesis defects [25]. On the other hand, parasites with a motility defect, albeit without impaired cell division were shown to be infective in mice, leaving the question of in vivo relevance of

Fig. 6. Different life cycle stages of Trypanosoma brucei reveal trypomastigote morphology. The anterior (A) and posterior (P) cell poles are indicated.

directional motility for long term survival open for studies in more natural infection models [79]. Since the first quantitative descriptions of T. brucei, the BSFs have been classified as either long and slender, or short and stumpy – albeit with a continuous range of intermediate forms [80] (Fig. 6). At the same time, a persistent fluctuation between the two morphotypes in the host has been observed. Host parasitemia rises when the long slender BSFs proliferate, and falls when the host’s adaptive immune response begins to eliminate the parasites. BSFs have a dense surface coat composed of a single isoform of densely packed variant surface glycoprotein (VSG); at any time, single trypanosomes can change their coat in a process of antigenic variation. This allows these cells and their clonal offspring to undergo another period of unchecked proliferation while the immune system has to generate new antibodies [81]. With each wave of rising parasitemia, a quorum sensing mechanism is activated by a trypanosomederived “stumpy induction factor” (SIF). SIF induces differentiation of BSF into short stumpy forms [82]. The stumpy cells accumulate because they are less susceptible to the developing antibody response [83]. They are cell cycle-arrested and constitute the tsetse fly infective form. In this way, the parasite balances the requirements for its own and its host’s survival with the need for dispersal by switching hosts [84]. Besides surviving whilst being swept through the mammalian circulation, the BSF trypanosomes can also leave the lymph system and blood vessels, invade tissue spaces [85,86], and ultimately the central nervous system (CNS). Once in the CNS, they cause the neurological symptoms that give sleeping sickness its name [87,88]. Throughout an infection, the parasites spur a multitude of immunological interactions between themselves and the host which control the progress of disease [89]. While the parasite’s long term survival is enabled by antigenic variation, i.e. switching to the expression of a different VSG, the cells also have the capacity to directly eliminate VSG-bound antibodies using endocytosis [83]. This strategy is effective when trypanosomes swim directionally in the anterior direction, allowing the hydrodynamic drag force of the surrounding fluid to push the surface-bound antibodies towards the cell

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

posterior. There, they are swept into the flagellar pocket and endocytosed [90]. The elucidation of this antibody clearance mechanism catalysed the detailed analysis of trypanosome motility. This effort was necessary in order to show how the parasites reach the swimming speeds in blood that are theoretically required by hydrodynamics for antibody clearance to be effective. The cells swimming speeds in culture medium turned out to be too low and the exact swimming mechanism of trypanosomes was unclear at the time [32]. T. brucei BSF swim forward by producing tip-to-base flagellar waves, but also produce occasional reverse waves, similar to the other kinetoplastids [31,91]. Due to the extensive helical attachment of the flagellum to the cell body in the trypomastigote form, the resulting movement is in a three-dimensional helical pattern. In low viscosity culture medium the flagellar wave direction often reverses, producing opposing forces that generate a characteristic tumbling motion without efficient translocation. When simulating the situation in blood or other highly viscous environments, the trypanosomes were shown to swim faster and more persistently, fulfilling the criteria for efficient antibody clearance [32]. Under the mechanical influence of obstacles or high viscosity, the trypanosomes can also exhibit sustained backward movement due to persistent base-to-tip flagellar beating [32]. This flexibility of swimming modes and the ability to respond to mechanical cues under conditions mimicking those in the bloodstream or tissue spaces probably plays an important role in manoeuvring between these highly variable environments throughout the life-cycle of the parasite. These characteristics have also been analysed in trypanosomes pathogenic to animals. They were found to be quite species-specific and correlate with the observed preference of different niches occupied in the mammalian hosts. (Bargul et al., in revision). Forward propulsion in T. brucei is dependent on the functionality of the outer axonemal dynein motors. Flagellar tip-to-base beats are impeded by defects in the outer dynein arms [91,92]. Mutants with these defects continue to produce flagellar waves from baseto-tip, which are able to propel the trypanosomes backwards in sufficiently viscous surroundings [90]. These motility phenotypes could be useful for the analysis of axonemal mechanisms and their control, but so far no mutant exclusively blocking backward movement has been described. An unusual feature of T. brucei is the simultaneous occurrence of forward and reverse waves [32]. This does not seem to occur in other trypanosomatids, where a beat reversal is only initiated after a completed wave [31], although a rare observation was recorded in the free flagellum of Crithidia oncopelti [93]. The simultaneous propagation of axonemal bending in opposing directions poses an interesting challenge in theoretical modelling of flagellar motility. The trypanosomes’ infiltration of the brain has been a longstanding enigma. The parasites can enter the CNS by crossing the blood–cerebrospinal fluid (CSF) barrier in the choroid plexus via fenestrated blood vessels, and possibly by opening tight junctions in the epithelial cell layer [86]. They repeatedly cross this barrier during waves of high parasitemia and may develop into a very long slender, fast, and directional swimming morphotype. This morphotype could be optimised for crossing CSF spaces, where they do not survive, and into safer microcompartments. From there, they have been reported to reappear in the blood, as well as moving on into the meningeal system and causing meningitis. If they are not controlled by the host, they eventually penetrate into the brain parenchyma, leading to the encephalitic final stage of the disease [87]. A recent study has imaged T. brucei in the cortical meninges of intact living mice brains, together with the accumulation of lymphocytes that occurs during meningitis [94]. Extravascular trypanosomes were observed to accumulate in the meningeal space following the accumulation of lymphocytes. They became less motile as the infection progressed and eventually disappeared. No extravascular parasites

7

were detected in the parenchyma or returning to the bloodstream. Conversely, a previous study, also using intra-vital microscopy of the mouse cortex, has shown early migration of trypanosomes into the parenchyma, suggesting the direct crossing of the blood–brainbarrier [95]. Until the BSF trypanosomes manage to invade the brain, their differentiation into the cell cycle-arrested, short stumpy form ensures that the vertebrate host is not killed by uncontrolled proliferation of the slender forms. Additionally, the stumpy form appears sturdier than slender forms and in several ways is adapted to survive and develop in the insect host [96–99]. Only stumpy cells can survive the environment in the fly gut, and differentiate within 24 h into procyclic forms. The procyclics migrate into the tissue of the fly gut and begin a complex developmental cycle [100]. It is noteworthy that the motility of individual life cycle stages differs greatly. The slender BSF forms, which are strongly motile, transform successively to the less motile stumpy forms and then to the very motile procyclic trypomastigotes (Fig. 6). Nothing is known about the role(s) of the varying motility, although reduced persistent swimming has been shown to impair infection of the tsetse fly [101] Further development in the fly is characterised by a succession of several morphotypes, including the transition from trypomastigote to epimastigote stages and back again. Whereas the morphological changes are relatively well described, there is currently only speculation about the role of the parasite’s motility in the traversal of the diverse environmental surroundings of the fly’s organs and the survival of single cells at developmental bottlenecks [74,100,102–104]. Though the characterisation of the trypanosomes’ physical environment in the insect host awaits analysis, it has meanwhile been shown that cultured procyclic cells exhibit a form of collective behaviour called “social motility” when allowed to grow on agarose plates [105]. The procyclics are able to proliferate in a thin fluid film on agar surfaces and appear to exhibit radiating patterns of movement outwards from the original colonies. This behaviour has been shown to be specific for an early form of procyclic cells and thus is thought to mimic an early step in the colonisation of the tsetse’s midgut, where the parasite multiplies to great densities [106]. Live imaging with single cell resolution in the tsetse fly to analyse such phenomena is demanding, but is currently being undertaken and should provide insight into the nature of the parasite’s impressive collective motion patterns. Complementing the emerging in vivo motility data, an alternative in silico approach has shown that by numerically modelling different trypanosome morphotypes in a realistic hydrodynamic environment, it is possible to generate morphological and motility data for any combination of cell type and physical environment, thus allowing predictions of motile behaviour in the natural host–parasite system [107]. Last but not least, T. brucei produces haploid gametes for sexual reproduction in the salivary glands of the tsetse fly [108]. The meiotic stages manifest as epimastigote cells in early salivary gland infections. They represent a normal stage of development, between attached epimastigotes and developing metacyclics [109]. The resulting gametes were observed in vitro and represent the only T. brucei promastigote-like stage, with long free flagella that intertwine when two gametes meet for fusion [108]. It would be interesting to detail the gametes’ motion and recognition behaviour in the natural environment of the salivary glands. All kinetoplastid flagella have a highly organised lattice-like structure bound tightly to the length of the axoneme, called the paraflagellar rod (PFR) [110]. The complete structure is surrounded by the flagellar membrane, which is attached in a defined, complex order to the cell body membrane in trypomastigote cells [111], and thus inseparable from flagellar motility. Despite substantial knowledge of the composition and structure of the PFR, as well as of distinct detrimental effects on flagellar motility in several PFR

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model

ARTICLE IN PRESS

YSCDB-1865; No. of Pages 15

T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

8

Fig. 7. The ‘minimalistic morphology’ of the Plasmodium microgamete.

mutants, an exact mechanism in motility has not been shown. Many hypotheses exist on structural functions of the PFR, and potential roles in signalling are being added in motility, as well as in sensory flagellar functions [110,112]. 3.3. Apicomplexa – Plasmodium Although not a flagellate parasite during its asexual proliferative life cycle, the malaria pathogen Plasmodium deserves mention for its remarkable flagellate male microgamete. Despite its very brief appearance in the sexual life stage, it was famously responsible for the discovery of the parasite. Upon uptake by the mosquito vector (Anopheles), gametocytes from the blood of the human host convert to flagellated gametes in a matter of minutes. The spermlike male microgametes are motile and survive less than an hour in the insect’s midgut, where they fertilise the female gamete in an interesting and understudied process [113]. Although Plasmodium gametes have been biologically difficult to assess, they have recently been introduced as a unique, mathematically tractable model system in a detailed motility study [114]. The microgamete represents an extremely simple, sperm-like motile cell with a classic axonemal structure, amenable to study due to the lack of extracellular layers or a hydrodynamically important cell body (Fig. 7). Interestingly, the flagellated microgametes of Plasmodium, which have to move in the densely packed bloodmeal in the insect gut and on the surface of female gametes, also show forward and backward movement, reminiscent of the trypanosomatids. Although the corresponding flagellar wave patterns have not been sufficiently resolved, the forward movement was quantified and shown to be due to tip-to-base flagellar waves, which is unusual for spermatozoa [114]. 4. Physics of flagellated microswimmers Analysis of the lifestyles of parasitic flagellates (as detailed in the previous section and summarised in Table 1) makes it clear that cellular motility – while utilising common structural components – can be tailored to a wide variety of environments and biological niches. The motility of flagellated parasites must also be adapted dynamically to varying physical environments. The cells are able to produce a multitude of motile behaviours by bending one or a few filamentous appendages and generating propulsive forces against viscous fluids and surfaces. These are described by the resistive force theory [115], a mathematical description of flagellar propulsion developed after in-depth analysis of the spermatozoon of the sea urchin Psammechinus miliaris [116]. This hydrodynamic theory is able to predict velocities and trajectories of flagellum-driven microswimmers in fluids and near surfaces [17,117]. The long slender shape of the flagellum can be moved forward with the force of a bending wave running along its long axis and pushing against the surrounding fluid in the opposite direction. This is possible because the hydrodynamic friction is lower for the rod-shaped body moving in the direction of the long axis than perpendicular to it [17,118].

Friction is the only relevant force for the propulsion of microswimmers. For microbes, water is a very viscous medium. The ratio of inertial forces to viscous forces, called the Reynolds number, is extremely low. Without inertia, the drag force of the oscillating flagellum immediately converts into active movement of the microorganism. Likewise, any interruption of force production will immediately stop the cell’s locomotion. An important consequence of this situation is that a simple reversal of movement will bring the swimmer back to exactly the same position, which means that propulsive movement must be non-reciprocal [118,119]. This criterion is fulfilled in flagellates with a unidirectional travelling wave or the ciliary power/recovery stroke cycle. This hydrodynamic concept is straightforward for free flagella swimming in fluids, but also applies to swimming near surfaces, where the validity of resistive force theory has been tested by high precision tracking of sperm movements [117]. The propulsion of flagellate parasites should therefore also be adequately described by the hydrodynamic model, with the prerequisite for testing being a high spatiotemporal resolution recording of the flagellar movement and the cell’s translocation. Such analysis of flagellar beats have been performed, as mentioned above, for free flagella with Leishmania [31], attached single flagella with T. brucei [32] and multiple flagella with Giardia [48] and Trichomonas [40]. In the latter two (multiflagellate) cases, the hydrodynamics of each individual flagellum could be modelled on the basis of the resistive force theory. The analysis allowed the determination of the exact role of each appendage for the parasite’s motility, including quantitative data on the forces produced and thus, allowing inferences for the mechanism of host invasion. The case of T. brucei is complicated by the flagellum being attached to a large part of the cell body in a helical turn, causing the cell to rotate around its long axis when swimming directionally [32]. Nevertheless, the free part of the flagellum generates planar flagellar waves, which produce the drag force to move the cell, usually forwards with a tip-to-base wave pulling the cell along. For directionally moving cells, flagellar force production has been measured to be in the low (1–4) pN range [120], which is similar to the range of forces measured for other single eukaryotic flagella [40]. The power needed for propulsion is quite low in comparison to the power actually produced by the flagellates [120], meaning the efficiency of the propulsion system is low. Calculations showed this generally to be the case for biological microswimmers and the efficiency was famously commented on as being an irrelevant item in the “bugs” energy budget, with their swimming likened to driving an economy car in a place of unlimited fuel supply [119]. For trypanosomes in the bloodstream, for example, there is an endless supply of glucose to fuel the parasites’ incessant motility. The biological significance of flagellar motility in the fast flowing habitat of the bloodstream is not immediately clear, but the ability to produce hydrodynamic flow forces along the cell surface by fast directional swimming – facilitating the removal of host antibodies – is one important function [90]. The trypanosomes are accordingly adapted to the conditions in the bloodstream and exhibit higher velocities and swimming persistence in blood or fluids of similar viscosity [32]. The in vivo behaviour of parasites can thus be significantly different from that observed under culture conditions. Another obvious but little studied physical aspect is the influence of the flow in the bloodstream on motility [121]. Even in sperm, the best analysed model of flagellar motility, the need to focus on the role of environmental viscosity and flow is recognised [122]. The trypanosomatids have to be adapted to more than one viscous fluid environment and cross several solid boundaries in order to successfully complete their infection cycles. They achieve this by

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

9

Table 1 Brief overview of aspects of the parasites’ flagellate stages and their relevance to human infection. Some species used in cited studies are listed, including zoonotic subspecies. Lengths refer to the whole cell including flagella. Human parasite

Human disease

Human transmission

Flagellate morphotypes

Role in infection

Trichomonas vaginalis

Trichomoniasis

Sexually transmitted disease

Trophozoite (4–32 ␮m)

Zoon. model: Tritrichomanas foetus

Animal disease: Bovine Trichomoniasis

Flagellates exchanged with urogenital fluids

4 anterior Fl. 1 attached Fl. forming undulating membrane

Motile in urogenital tract Transformation into amoeboid upon contact with vaginal epithelial cells and erosion of epithelium

Giardia lamblia syn. intestinalis syn. duodenalis

Giardiasis

Fecal-oral

Trophozoite (10–20 ␮m)

Leishmania spp.

Aflagellate cysts ingested with contaminated water or food Leishmaniasis

e.g. L. major L. donovani Trypanosoma cruzi

Chagas

4 lateral Fl. 2 ventral Fl. 2 caudal Fl.

Sand flies (>90 species)

Promastigote (15–30 ␮m)

Flagellates injected into the skin with saliva during blood meal

1 free anterior Fl.

Hemato-phagous bugs (Reduviidae)

Trypomastigote (15–24 ␮m)

Trypomastigote metacyclics deposited in the feeding wound with excretions

Epimastigote (15–24 ␮m) 1 attached flagellum with free anterior part

Sleeping sickness

T. b. gambiense

Zoon. model: T. b. brucei

Tsetse fly (Glossina sp.) Trypomastigotemetacyclics injected into the skin with saliva during blood meal

Animal disease: Nagana

(Salivarian trypanosomes)

Motile in insect Motility-dependent phagocytosis by macrophages in the skin. Intracellular transformation into the amastigote form, which proliferates and spreads to cause species-specific tissue pathology Trypomastigote motile in lymph and bloodstream. Phagocytosed by macrophages. Intracellular transformation to amastigote form, proliferation and transformation back to trypomastigote form, renewed dispersal and invasion of host cells Epimastigote motile in insect

(Stercorian trypanosomes) Trypanosoma bruceirhodesiense

Motile in small intestine Attachment to intestinal epithelium, proliferation and destruction of epithelial barrier

Trypomastigote in bloodstream slender form (23–33 ␮m) stumpy form (17–22 ␮m) Various trypo- and epimastigote forms during development in the tsetse fly (7–46 ␮m)

Trypomastigote bloodstream forms permanently extracellular and motile in lymph, bloodstream and tissue spaces, causing periodic fever Invasion of CNS eventually disrupts the sleep-wake cycle Insects forms permanently motile, free swimming or attached during complex development

1 attached flagellum with free anterior part

changing their morphology in distinctive ways [51]. These changes affect motile behaviour, as the cell body is deformed with the flagellar oscillations and the movement of the parasite depends on the topology of the flagellar attachment, as well as the morphology and flexibility of the cell body [123]. Based on high resolution video microscopy data, the complex swimming mechanism of BSF T. brucei was confirmed by a full scale hydrodynamic computer simulation, using multi-particle collision dynamics [107]. This simulation method was then used to test different trypanosomal morphotypes in silico. The approach not only allowed comparison to in vivo data when available, but also prediction of adaptive values of morphologies that have not yet been analysed in vivo or in culture. Simulations like these can be conveniently extended to test the influence of different viscosities, confining environments, hydrodynamic flow and collective behaviour of other flagellate parasites. These are all very active current topics in the research fields of microswimmers [17], but hardly anything is known about their biological relevance for host invasion and pathogenicity. In fact, studies of the other parasites have lagged behind T. brucei.

5. Technical perspective and challenges 5.1. Spatiotemporal resolution In order to fully understand flagellate parasites, integrative approaches are needed, bringing technology and research strategies from various disciplines together. One basic requirement for analysis of the flagellate motility mechanisms is high spatiotemporal microscopic resolution. Flagellates swim with velocities of tens of ␮m/s while producing oscillations of tens of Hz. The first sufficiently high-speed photograph series were recorded more than 50 years ago, when cameras were developed which attained around 500 frames per second (fps). Flagella and cilia from several organisms were recorded, amongst them apparently several Trypanosomatidae, although not all were published [93]. An important result obtained from resolving the flagellar waves was that they did not only run from base-to-tip, as in the case of sperm. Today, digital high-speed cameras are available that allow high resolution recordings with equivalent speeds. Although high-speed digital cameras and microscopy systems have been used since the

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15 10

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

end of the last century, setups able to record with sample rates above the standard 30 fps video rate were uncommon [124–126]. The tremendous advances in microscopy since then were focused on the improvement of resolution and sensitivity for sophisticated fluorescent detection methods. Super-resolution techniques are currently being developed to observe live cell dynamics with sufficient speed, down to the single-molecule level [127–129]. In addition to a wide spectrum of available commercial microscope systems for any level of resolution and budget available today, there is camera technology forthcoming which provides the speed and sensitivity sufficient for high-speed recording of biological processes in standard light and fluorescence microscopes. Many imaging applications can benefit from well-established techniques, performed with affordable yet state-of-the-art digital cameras [130,131]. Using high-speed microscopic imaging, human flagellate parasites have been recorded at frame rates from 150 to 500 fps in order to resolve single waves of flagella beating with 5–30 Hz. Note that the high sampling rates need not be that high in order to resolve the flagellar beat, but as the speeds of the waves travelling along the flagella reach hundreds of ␮m/s, they are necessary to determine exact wave shapes and deviations from such. By imaging with sufficient spatiotemporal resolution, several important mechanisms have been elucidated or suggested. The precise beating modes and propulsive contributions of single flagella in Trichomonas [40] and Giardia [48] were analysed and quantitatively modelled, enabling explanation of the precise swimming and steering of the parasites and thus important capabilities for invasion of the host intestinal tract. The flagellar motility of Leishmania was precisely determined [31] and could be interesting to (re)analyse in light of the importance of motility in host cell invasion [56]. The mechanism of T. brucei motility was elucidated, explaining the adaptation of the parasites’ motility to their host’s bloodstream [32]. The forward movement of Plasmodium microgametes has been quantified and was speculated to be adapted to the habitat in blood, analogous to trypanosomes [114]. 5.2. The third dimension A caveat with all currently available high-speed data is that it is two-dimensional. It remains extremely challenging to perform millisecond timescale volumetric imaging [132]. Two-dimensional data is incomplete and notoriously prone to misinterpretation when inferring three-dimensional movement, even when temporal resolution is sufficient. For example, in an analysis of T. brucei, the three-dimensional rotational movement resulting from the waves travelling along the flagellum attached in a helical pitch around the cell body was interpreted as active flagellar movement. This interpretation gave rise to a model of wave propagation in which the two ends of the flagellum produce helical waves of opposing chirality (bihelical motility). The helical waves were thought to change chirality periodically, which apparently resulted in cells ends flipping from side to side [133]. Two-dimensional data is not sufficient for such interpretations, as a unidirectional rotation can produce indistinguishable visual effects. The complex three-dimensional movement of trypanosomes was elaborately documented by several indirect methods, including a time-dependent tomography method, taking into account the exact cell morphology [32] and corroborated by numerical simulation [107]. The detailed analysis of the tip-to-base and base-to-tip waveforms explains in a physically plausible way how the cells move forwards, backwards, and tumble, depending on the frequencies of the reversing flagellar waves. Importantly, it does so without any need to invoke bilhelical motion. In order to record the three-dimensional motility of microswimmers directly, digital holographic microscopy methods have been

implemented [134,135]. Holography allows the non-invasive, rapid acquisition of the complete optic field in a three dimensional volume. Objects swimming unconstrained in this volume are illuminated with a coherent light source and their diffraction patterns are digitally captured. This data contains all the information needed to numerically calculate the three-dimensional position and size of objects in the optic field. The technology allows for high throughput tracking of numerous, fast microscopic objects and is thus being developed further in order to quantitatively analyse the motility of bacteria [136] and sperm [137]. The holographic analyses of T. brucei populations have produced data on motility patterns of different trypanosome strains and their temperature dependence, as well as the effect of motility mutants and environmental confinement [138]. These results were complemented and explained by the high resolution data described above [32], resulting in a broad understanding of trypanosome motility. Combining both levels of analysis could allow the unravelling of genetic and physical components of flagellar motility. Low spatial resolution is a drawback of holographic techniques, but in an analysis of the Plasmodium microgamete, it was possible to calculate three-dimensional waveform data from high-speed recordings. This showed a lack of chirality bias in the unconstrained axoneme [114]. 5.3. Tissue models A transition from two- to three-dimensional analysis is also underway in cell culture techniques. Realising the need to reproduce relevant physiological behaviour in cells, rapid progress is being made in establishing biomimetic tissue models. Cells intimately interact with extracellular matrices and interstitial fluids in heterogenous micro-environments in vivo. These conditions can be partially reproduced by co-culturing cells in three-dimensional materials and integrating microfluidic techniques into the tissue models. Microfluidics allow for perfusion of tissues, in order to control media exchange and signal gradients and should eventually allow vascularisation [139]. Exciting progress is being made in producing complete organ-on-a-chip microfluidic bioassays [140,141], and also assays for sperm motility [142]. Finally, the abovementioned holographic microscopy methods are being combined with microfluidic systems to create compact, high-throughput, quantitative single cell motility assays [143,144]. The analysis of human flagellate parasites needs to be transferred to the level of tissue and organ interactions, in order to assess the complex mechanisms of infection and pathogenesis. The role of flagellar motility has started to be recognised in several important stages of the parasites’ life cycles, but so far, virtually all high resolution studies capable of addressing motility mechanisms have been carried out under in vitro cell culture conditions with strains adapted to axenic culture. It has been well-documented that in trypanosomatids there can be significant differences in morphology and behaviour between strains. In particular, changes occur during long term in vitro cultivation of T. brucei populations, including loss of infectivity [145]. In mammalian cell culture studies with Leishmania and T. cruzi various strains and model systems exhibit important differences in behaviour and infectivity [56,146]. Environmental changes play an important role in the parasites’ development from one morphotype into another and their journey across tissue and host boundaries. These changes need to be investigated in defined and tractable tissue and organ models. 5.4. In vivo analysis Parasites ultimately need to be studied in the natural environments of their hosts. For human flagellate parasites, this means either obtaining biopsies [147,148], which could be used for

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

ex vivo motility analyses, or relying on mammalian (predominantly rodent) models for in vivo analyses [149–152]. The Plasmodium research community has led the way in this regard, making early use of in vivo imaging techniques to investigate the infection routes of parasites [153,154]. Bioluminescence imaging of trypanosomatids has recently been used to monitor the dynamics of mouse infections in real time. The genetic reporters required to make the parasites luminescent have been improved, making the detection of low cell numbers possible and thus the study of tissue tropism and pathogenesis during chronic infections is now feasible [155–160]. Intra-vital microscopy methods depend on the optical accessibility of the parasite in the host tissue, whether in an artificial system or a living animal. The inability to optically penetrate a thick three-dimensional environment usually precludes standard microscopy techniques. Two-photon microscopy allows the recording of fluorescent signals up to around one millimetre tissue depth, and features other advantages for in vivo imaging, in particular low phototoxicity [161]. It has been used extensively for imaging cellular motility in the immune system and the dynamic host–pathogen interactions in diverse tissues [162,163]. Infection processes have been visualised in the skin [164], especially with Leishmania. After injection, the relatively immobile parasite was shown to be phagocytosed by rapidly mobilised neutrophils accumulating at the wound site [165]. T. brucei has been imaged in the peripheral cerebral cortex through the thinned skull of living mice [94]. Previous imaging of trypanosomes in the mouse cortex was carried out with a single photon confocal microscope, necessitating surgery to access the peripheral cerebral tissue [95]. These studies have allowed the first in vivo glimpses into the pathogenic invasion of the nervous system, while leaving numerous questions to answer. Of relevance to the future analysis of intestinal parasites, challenging methodology has recently been demonstrated for intra-vital two-photon imaging of the gastrointestinal tract [166]. A major challenge in studies of this kind, is the immobilisation of the live animal’s tissue, without compromising structure and functionality. With the low acquisition speeds of two-photon microscopy, any spatial perturbation in the living system to be observed is detrimental to high resolution imaging. The challenge of high spatiotemporal resolution for the analysis of motile parasites (see Section 5.1), necessitates a trade-off between acquisition time and imaging penetration depth. For near-surface microscopy, alternatives like spinning–disk confocal microscopy have been successfully used, e.g. in imaging the infection of Plasmodium in vivo [167] and are constantly being improved [168]. 5.5. Molecular genetic tools The combination of dedicated observation techniques and diverse analytic approaches from physical, computational and life science disciplines is necessary to generate a holistic understanding of parasite behaviour during host infection. A prerequisite for interpreting biological function is the generation and comparative analysis of genetic phenotypes. The genomes of the human flagellar parasites are completely accessible via EuPathDB [169], and molecular genetic tools are available to varying degrees in different parasites [170,171]. The trypanosomatid research community have led the development of genetic expression and manipulation systems including RNAi, especially for T. brucei [172,173]. Recently, genome editing using the CRISPR-Cas9 system has been applied to parasites for which important genetic tools are unavailable or ineffective, i.e. T. cruzi [174] and Leishmania [175]. Appropriately, the first targets of these new tools were genes whose depletion produces motility effects, i.e. PFR components. When these proteins are depleted in trypanosomatids, the cells are viable but manifest evident motility phenotypes [176–178]. Motility phenotypes are however difficult to interpret, due to the functional pleiotropy of

11

the flagellum, which complicates the distinction between direct, indirect, and stage-specific effects on the cell cycle, morphogenesis, cytokinesis and host invasion [25,112]. The stage should now be set to technically and functionally resolve and explain the flagellar motility of our parasites, which continues to fascinate, as it has ever since Leeuwenhoek’s first observations of his very own “animalcules a-moving very prettily” [179].

Acknowledgements We are grateful to Brooke Morriswood for his input and critical reading of the manuscript. We also thank Ines Subota and Sarah Schuster for valuable discussion. Many thanks to the reviewers for excellent suggestions. Our work is supported by the DFG Priority Programme “Microswimmers” (SPP 1726) and the DFG “GermanAfrican Cooperation Projects in Infectology” (PAK 296). We wish to apologise to the many researchers whose work we were not able to cite directly owing to space constraints and the scope of the review.

References [1] Mitchell DR. The evolution of eukaryotic cilia and flagella as motile and sensory organelles. Adv. Exp. Med. Biol 2007;607:130–40, http://dx.doi.org/10.1007/978-0-387-74021-8 11. [2] He D, Fiz-Palacios O, Fu C-J, Fehling J, Tsai C-C, Baldauf SL. An alternative root for the eukaryote tree of life. Curr. Biol 2014;24:465–70, http://dx.doi.org/10.1016/j.cub.2014.01.036. [3] Adl SM, Simpson AGB, Lane CE, Lukeˇs J, Bass D, Bowser SS, et al. The revised classification of eukaryotes. J. Eukaryot. Microbiol 2012;59:429–514, http://dx.doi.org/10.1111/j.1550-7408.2012.00644.x. [4] de Vargas C, Audic S, Henry N, Decelle J, Mahé F, Logares R, et al. Eukaryotic plankton diversity in the sunlit ocean. Science 2015;348:1261605, http://dx.doi.org/10.1126/science.1261605. [5] Leadbeater BSC, McCready SMM. The flagellates; historical perspectives. In: Leadbeater BSC, Green JC, editors. Flagellates Unity Divers. Evol. New York: Taylor & Francis, London; 2000. [6] Cavalier-Smith T. The excavate protozoan phyla Metamonada Grassé emend. (Anaeromonadea, Parabasalia, Carpediemonas, Eopharyngia) and Loukozoa emend. (Jakobea, Malawimonas): their evolutionary affinities and new higher taxa. Int. J. Syst. Evol. Microbiol 2003;53:1741–58. ´ c J, Votypka ´ [7] Lukeˇs J, Skalicky´ T, Tyˇ J, Yurchenko V. Evolution of parasitism in kinetoplastid flagellates. Mol. Biochem. Parasitol 2014;195:115–22, http://dx.doi.org/10.1016/j.molbiopara.2014.05.007. [8] Heintzelman M. Gliding motility in apicomplexan parasites; 2015. [9] Martinez AJ, Visvesvara GS. Free-living, amphizoic and opportunistic amebas. Brain Pathol. Zurich Switz 1997;7:583–98. [10] Marciano-Cabral F, Cabral GA. The immune response to Naegleria fowleri amebae and pathogenesis of infection. FEMS Immunol. Med. Microbiol 2007;51:243–59, http://dx.doi.org/10.1111/j.1574-695X.2007.00332.x. [11] Fritz-Laylin LK, Assaf ZJ, Chen S, Cande WZ. Naegleria gruberi de novo basal body assembly occurs via stepwise incorporation of conserved proteins. Eukaryot. Cell 2010;9:860–5, http://dx.doi.org/10.1128/EC.00381-09. [12] Jeuck A, Arndt H. A short guide to common heterotrophic flagellates of freshwater habitats based on the morphology of living organisms. Protist 2013;164:842–60, http://dx.doi.org/10.1016/j.protis.2013.08.003. [13] Raven JA. The flagellate condition. In: Leadbeater BSC, Green JC, editors. Flagellates Unity Divers. Evol. New York: Taylor & Francis, London; 2000. [14] Matthews KR. Controlling and coordinating development in vector-transmitted parasites. Science 2011;331:1149–53, http://dx.doi.org/10.1126/science.1198077. [15] Seco-Hidalgo V, Osuna A, Pablos LMD. To bet or not to bet: deciphering cell to cell variation in protozoan infections. Trends Parasitol 2015;31:350–6, http://dx.doi.org/10.1016/j.pt.2015.05.004. [16] Dheilly NM, Poulin R, Thomas F. Biological warfare: microorganisms as drivers of host–parasite interactions. Infect. Genet. Evol 2015;34:251–9, http://dx.doi.org/10.1016/j.meegid.2015.05.027. [17] Elgeti J, Winkler RG, Gompper G. Physics of microswimmers—single particle motion and collective behavior: a review. Rep. Prog. Phys 2015;78:056601, http://dx.doi.org/10.1088/0034-4885/78/5/056601. [18] Lindemann CB, Lesich KA. The geometric clutch at 20: stripping gears or gaining traction? Reproduction 2015;150:R45–53, http://dx.doi.org/10.1530/REP-14-0498. [19] Lindemann CB, Lesich KA. Flagellar and ciliary beating: the proven and the possible. J. Cell Sci 2010;123:519–28, http://dx.doi.org/10.1242/jcs.051326.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15 12

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

[20] Woolley DM. Flagellar oscillation: a commentary on proposed mechanisms. Biol. Rev 2010;85:453–70, http://dx.doi.org/10.1111/j.1469-185X.2009.00110.x. [21] Brokaw CJ. Thinking about flagellar oscillation. Cell Motil. Cytoskeleton 2009;66:425–36, http://dx.doi.org/10.1002/cm.20313. [22] Diniz MC, Pacheco ACL, Farias KM, de Oliveira DM. The eukaryotic flagellum makes the day: novel and unforeseen roles uncovered after post-genomics and proteomics data. Curr. Protein Pept. Sci 2012;13:524–46, http://dx.doi.org/10.2174/138920312803582951. [23] Pazour GJ, Agrin N, Leszyk J, Witman GB. Proteomic analysis of a eukaryotic cilium. J. Cell Biol 2005;170:103–13, http://dx.doi.org/10.1083/jcb.200504008. [24] Baker MA, Naumovski N, Hetherington L, Weinberg A, Velkov T, Aitken RJ. Head and flagella subcompartmental proteomic analysis of human spermatozoa. Proteomics 2013;13:61–74, http://dx.doi.org/10.1002/pmic.201200350. [25] Broadhead R, Dawe HR, Farr H, Griffiths S, Hart SR, Portman N, et al. Flagellar motility is required for the viability of the bloodstream trypanosome. Nature 2006;440:224–7, http://dx.doi.org/10.1038/nature04541. [26] Oberholzer M, Langousis G, Nguyen HT, Saada EA, Shimogawa MM, Jonsson ZO, et al. Independent analysis of the flagellum surface and matrix proteomes provides insight into flagellum signaling in mammalian-infectious Trypanosoma brucei. Mol. Cell. Proteomics 2011;10:M111, http://dx.doi.org/10.1074/mcp.M111.010538, 010538. [27] Subota I, Julkowska D, Vincensini L, Reeg N, Buisson J, Blisnick T, et al. Proteomic analysis of intact flagella of procyclic Trypanosoma brucei cells identifies novel flagellar proteins with unique sub-localization and dynamics. Mol. Cell. Proteomics 2014;13:1769–86, http://dx.doi.org/10.1074/mcp.M113.033357. [28] Brooks ER, Wallingford JB. Multiciliated cells. Curr. Biol 2014;24:R973–82, http://dx.doi.org/10.1016/j.cub.2014.08.047. [29] Wan KY, Leptos KC, Goldstein RE. Lag, lock, sync, slip: the many “phases” of coupled flagella. J. R. Soc. Interface 2014;11:20131160, http://dx.doi.org/10.1098/rsif.2013.1160. [30] Hyams JS, Borisy GG. Isolated flagellar apparatus of Chlamydomonas: characterization of forward swimming and alteration of waveform and reversal of motion by calcium ions in vitro. J. Cell Sci 1978;33:235–53. [31] Gadelha C, Wickstead B, Gull K. Flagellar and ciliary beating in trypanosome motility. Cell Motil. Cytoskeleton 2007;64:629–43, http://dx.doi.org/10.1002/cm.20210. [32] Heddergott N, Krüger T, Babu SB, Wei A, Stellamanns E, Uppaluri S, et al. Trypanosome motion represents an adaptation to the crowded environment of the vertebrate bloodstream. PLoS Pathog 2012;8:e1003023, http://dx.doi.org/10.1371/journal.ppat.1003023. [33] Maritz JM, Land KM, Carlton JM, Hirt RP. What is the importance of zoonotic trichomonads for human health? Trends Parasitol 2014;30:333–41, http://dx.doi.org/10.1016/j.pt.2014.05.005. [34] Poole DN, McClelland RS. Global epidemiology of Trichomonas vaginalis. Sex. Transm. Infect 2013;89:418–22, http://dx.doi.org/10.1136/sextrans-2013-051075. [35] Brugerolle G. Flagellar and cytoskeletal systems in amitochondrial flagellates: Archamoeba, Metamonada and Parabasala. Protoplasma 1991;164:70–90, http://dx.doi.org/10.1007/BF01320816. [36] de Andrade Rosa I, de Souza W, Benchimol M. High-resolution scanning electron microscopy of the cytoskeleton of Tritrichomonas foetus. J. Struct. Biol 2013;183:412–8, http://dx.doi.org/10.1016/j.jsb.2013.07.002. [37] Kusdian G, Woehle C, Martin WF, Gould SB. The actin-based machinery of Trichomonas vaginalis mediates flagellate-amoeboid transition and migration across host tissue. Cell. Microbiol 2013;15:1707–21, http://dx.doi.org/10.1111/cmi.12144. [38] Kusdian G, Gould SB. The biology of Trichomonas vaginalis in the light of urogenital tract infection. Mol. Biochem. Parasitol 2014;198:92–9, http://dx.doi.org/10.1016/j.molbiopara.2015.01.004. [39] Hirt RP. Trichomonas vaginalis virulence factors: an integrative overview. Sex. Transm. Infect 2013;89:439–43, http://dx.doi.org/10.1136/sextrans-2013-051105. [40] Lenaghan SC, Nwandu-Vincent S, Reese BE, Zhang M. Unlocking the secrets of multi-flagellated propulsion: drawing insights from Tritrichomonas foetus. J. R. Soc. Interface 2014;11:20131149, http://dx.doi.org/10.1098/rsif.2013.1149. [41] Monteiro-Leal LH, Farina M, Benchimol M, Kachar B, De Souza W. Coordinated flagellar and ciliary beating in the protozoon Tritrichomonas foetus. J. Eukaryot. Microbiol 1995;42:709–14, http://dx.doi.org/10.1111/j.1550-7408.1995.tb01621.x. [42] Monteiro-Leal LH, Farina M, de Souza W. Free movement of Tritrichomonas foetus in a liquid medium: a video-microscopy study. Cell Motil. Cytoskeleton 1996;34:206–14, http://dx.doi.org/10.1002/ (SICI)1097-0169(1996)34:3<206::AID-CM4>3.0.CO;2-7. [43] Bartelt LA, Sartor RB. Advances in understanding Giardia: determinants and mechanisms of chronic sequelae. F1000Prime Rep 2015;7, http://dx.doi.org/10.12703/P7-62.

[44] House SA, Richter DJ, Pham JK, Dawson SC. Giardia flagellar motility is not directly required to maintain attachment to surfaces. PLoS Pathog 2011;7:e1002167, http://dx.doi.org/10.1371/journal.ppat.1002167. [45] Dawson SC, House SA. Life with eight flagella: flagellar assembly and division in Giardia. Curr. Opin. Microbiol 2010;13:480–90, http://dx.doi.org/10.1016/j.mib.2010.05.014. [46] Carvalho KP, Monteiro-Leal LH. The caudal complex of Giardia lamblia and its relation to motility. Exp. Parasitol 2004;108:154–62, http://dx.doi.org/10.1016/j.exppara.2004.08.007. [47] Lenaghan SC, Davis CA, Henson WR, Zhang Z, Zhang M. High-speed microscopic imaging of flagella motility and swimming in Giardia lamblia trophozoites. Proc. Natl. Acad. Sci. U. S. A 2011;108:E550–8, http://dx.doi.org/10.1073/pnas.1106904108. [48] Lenaghan SC, Chen J, Zhang M. Modeling and analysis of propulsion in the multiflagellated micoorganism Giardia lamblia. Phys. Rev. E: Stat. Nonlin. Soft Matter Phys 2013;88:012726. [49] Peyer KE, Zhang L, Nelson BJ. Bio-inspired magnetic swimming microrobots for biomedical applications. Nanoscale 2013;5:1259–72, http://dx.doi.org/10.1039/c2nr32554c. ´ [50] Maslov DA, Votypka J, Yurchenko V, Lukeˇs J. Diversity and phylogeny of insect trypanosomatids: all that is hidden shall be revealed. Trends Parasitol 2013;29:43–52, http://dx.doi.org/10.1016/j.pt.2012.11.001. [51] Wheeler RJ, Gluenz E, Gull K. The limits on trypanosomatid morphological diversity. PLoS ONE 2013;8:e79581, http://dx.doi.org/10.1371/journal.pone.0079581. [52] Bates PA. Leishmania sand fly interaction: progress and challenges. Curr. Opin. Microbiol 2008;11:340–4, http://dx.doi.org/10.1016/j.mib.2008.06.003. [53] Adhiambo C, Forney JD, Asai DJ, LeBowitz JH. The two cytoplasmic dynein-2 isoforms in Leishmania mexicana perform separate functions. Mol. Biochem. Parasitol 2005;143:216–25, http://dx.doi.org/10.1016/j.molbiopara.2005.04.017. [54] Cuvillier A, Miranda JC, Ambit A, Barral A, Merlin G. Abortive infection of Lutzomyia longipalpis insect vectors by aflagellated LdARL-3A-Q70L overexpressing Leishmania amazonensis parasites. Cell. Microbiol 2003;5:717–28. [55] Barros VC, Oliveira JS, Melo MN, Gontijo NF. Leishmania amazonensis: chemotaxic and osmotaxic responses in promastigotes and their probable role in development in the phlebotomine gut. Exp. Parasitol 2006;112:152–7, http://dx.doi.org/10.1016/j.exppara.2005.10.005. [56] Forestier C-L, Machu C, Loussert C, Pescher P, Späth GF. Imaging host cell-Leishmania interaction dynamics implicates parasite motility, lysosome recruitment, and host cell wounding in the infection process. Cell Host Microbe 2011;9:319–30, http://dx.doi.org/10.1016/j.chom.2011.03.011. [57] Alexander J. Unusual axonemal doublet arrangements in the flagellum of Leishmania amastigotes. Trans. R. Soc. Trop. Med. Hyg 1978;72:345–7. [58] Simpson CF, Harvey JW, French TW. Ultrastructure of amastigotes of Leishmania donovani in the bone marrow of a dog. Am. J. Vet. Res 1982;43:1684–6. [59] Gluenz E, Höög JL, Smith AE, Dawe HR, Shaw MK, Gull K. Beyond 9+0: noncanonical axoneme structures characterize sensory cilia from protists to humans. FASEB J 2010;24:3117–21, http://dx.doi.org/10.1096/fj.09-151381. [60] Bartholomeu DC, de Paiva RMC, Mendes TAO, DaRocha WD, Teixeira SMR. Unveiling the intracellular survival gene kit of trypanosomatid parasites. PLoS Pathog 2014;10:e1004399, http://dx.doi.org/10.1371/journal.ppat.1004399. [61] de Morais CGV, Castro Lima AK, Terra R, dos Santos RF, Da-Silva SA, Dutra PM, et al. The dialogue of the host–parasite relationship: Leishmania spp. and Trypanosoma cruzi infection. BioMed Res. Int 2015;2015:e324915, http://dx.doi.org/10.1155/2015/324915. [62] Ballesteros-Rodea G, Santillán M, Martínez-Calvillo S, Manning-Cela R. Flagellar motility of Trypanosoma cruzi epimastigotes. J. Biomed. Biotechnol 2012;2012:520380, http://dx.doi.org/10.1155/2012/520380. [63] Walker DM, Oghumu S, Gupta G, McGwire BS, Drew ME, Satoskar AR. Mechanisms of cellular invasion by intracellular parasites. Cell. Mol. Life Sci. CMLS 2014;71:1245–63, http://dx.doi.org/10.1007/s00018-013-1491-1. [64] Johnson CA, Rachakonda G, Kleshchenko YY, Nde PN, Madison MN, Pratap S, et al. Cellular response to Trypanosoma cruzi infection induces secretion of defensin ␣-1, which damages the flagellum, neutralizes trypanosome motility, and inhibits infection. Infect. Immun 2013;81:4139–48, http://dx.doi.org/10.1128/IAI.01459-12. [65] Andrade LO, Andrews NW. The Trypanosoma cruzi – host-cell interplay: location, invasion, retention. Nat. Rev. Microbiol 2005;3:819–23, http://dx.doi.org/10.1038/nrmicro1249. [66] Zingales B, Miles MA, Campbell DA, Tibayrenc M, Macedo AM, Teixeira MMG, et al. The revised Trypanosoma cruzi subspecific nomenclature: rationale, epidemiological relevance and research applications. Infect. Genet. Evol 2012;12:240–53, http://dx.doi.org/10.1016/j.meegid.2011.12.009. [67] McCall L-I, McKerrow JH. Determinants of disease phenotype in trypanosomatid parasites. Trends Parasitol 2014;30:342–9, http://dx.doi.org/10.1016/j.pt.2014.05.001.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

[68] Perez CJ, Lymbery AJ, Thompson RCA. Chagas disease: the challenge of polyparasitism? Trends Parasitol 2014;30:176–82, http://dx.doi.org/10.1016/j.pt.2014.01.008. [69] Dias GBM, Gruendling AP, Araújo SM, Gomes ML, Toledo MJde O. Evolution of infection in mice inoculated by the oral route with different developmental forms of Trypanosoma cruzi I and II. Exp. Parasitol 2013;135:511–7, http://dx.doi.org/10.1016/j.exppara.2013.08.013. [70] Pays E, Vanhollebeke B, Uzureau P, Lecordier L, Pérez-Morga D. The molecular arms race between African trypanosomes and humans. Nat. Rev. Microbiol 2014;12:575–84, http://dx.doi.org/10.1038/nrmicro3298. [71] Capewell P, Cooper A, Clucas C, Weir W, Macleod A. A co-evolutionary arms race: trypanosomes shaping the human genome, humans shaping the trypanosome genome. Parasitology 2015;142:S108–19, http://dx.doi.org/10.1017/S0031182014000602. [72] Matthews KR. 25 years of African trypanosome research: from description to molecular dissection and new drug discovery. Mol. Biochem. Parasitol 2015;200:30–40, http://dx.doi.org/10.1016/j.molbiopara.2015.01.006. [73] Vickerman K. Developmental cycles and biology of pathogenic trypanosomes. Br. Med. Bull 1985;41:105–14. [74] Rotureau B, Subota I, Buisson J, Bastin P. A new asymmetric division contributes to the continuous production of infective trypanosomes in the tsetse fly. Development 2012;139:1842–50, http://dx.doi.org/10.1242/dev.072611. [75] Abenga JN. A comparative pathology of Trypanosoma brucei infections. Glob. Adv. Res. J. Med. Med. Sci 2014;3:390–9. [76] Barry JD, Emery DL. Parasite development and host responses during the establishment of Trypanosoma brucei infection transmitted by tsetse fly. Parasitology 1984;88:67–84, http://dx.doi.org/10.1017/S0031182000054354. [77] Poltera AA. Pathology of human African trypanosomiasis with reference to experimental African trypanosomiasis and infections of the central nervous system. Br. Med. Bull 1985;41:169–74. [78] Malvy D, Chappuis F. Sleeping sickness. Clin. Microbiol. Infect 2011;17:986–95, http://dx.doi.org/10.1111/j.1469-0691.2011.03536.x. [79] Kisalu NK, Langousis G, Bentolila LA, Ralston KS, Hill KL. Mouse infection and pathogenesis by Trypanosoma brucei motility mutants. Cell. Microbiol 2014;16:912–24, http://dx.doi.org/10.1111/cmi.12244. [80] Bruce D, Hamerton AE, Bateman HR, Mackie FP. Trypanosome diseases of domestic animals in Uganda. II: Trypanosoma brucei (Plimmer and Bradford). Proc. R. Soc. Lond. B: Biol. Sci 1910;83:1–14, http://dx.doi.org/10.1098/rspb.1910.0062. [81] Mugnier MR, Cross GAM, Papavasiliou FN. The in vivo dynamics of antigenic variation in Trypanosoma brucei. Science 2015;347:1470–3, http://dx.doi.org/10.1126/science.aaa4502. [82] Mony BM, MacGregor P, Ivens A, Rojas F, Cowton A, Young J, et al. Genome-wide dissection of the quorum sensing signalling pathway in Trypanosoma brucei. Nature 2014;505:681–5, http://dx.doi.org/10.1038/nature12864. [83] McLintock LM, Turner CM, Vickerman K. Comparison of the effects of immune killing mechanisms on Trypanosoma brucei parasites of slender and stumpy morphology. Parasite Immunol 1993;15:475–80. [84] MacGregor P, Szö"or B, Savill NJ, Matthews KR. Trypanosomal immune evasion, chronicity and transmission: an elegant balancing act. Nat. Rev. Microbiol 2012;10:431–8, http://dx.doi.org/10.1038/nrmicro2779. [85] Losos GJ, Ikede BO. Review of pathology of diseases in domestic and laboratory animals caused by Trypanosoma congolense, T. vivax, T. brucei, T. rhodesiense and T. gambiense. Vet. Pathol. Online 1972;9:1–79, http://dx.doi.org/10.1177/030098587200901s01. [86] Mogk S, Meiwes A, Shtopel S, Schraermeyer U, Lazarus M, Kubata B, et al. Cyclical appearance of African trypanosomes in the cerebrospinal fluid: new insights in how trypanosomes enter the CNS. PLoS ONE 2014;9:e91372, http://dx.doi.org/10.1371/journal.pone.0091372. [87] Mogk S, Meiwes A, Boßelmann CM, Wolburg H, Duszenko M. The lane to the brain: how African trypanosomes invade the CNS. Trends Parasitol 2014;30:470–7, http://dx.doi.org/10.1016/j.pt.2014.08.002. [88] Ueno N, Lodoen MB. From the blood to the brain: avenues of eukaryotic pathogen dissemination to the central nervous system. Curr. Opin. Microbiol 2015;26:53–9, http://dx.doi.org/10.1016/j.mib.2015.05.006. [89] Beschin A, Van Den Abbeele J, De Baetselier P, Pays E. African trypanosome control in the insect vector and mammalian host. Trends Parasitol 2014;30:538–47, http://dx.doi.org/10.1016/j.pt.2014.08.006. [90] Engstler M, Pfohl T, Herminghaus S, Boshart M, Wiegertjes G, Heddergott N, et al. Hydrodynamic flow-mediated protein sorting on the cell surface of trypanosomes. Cell 2007;131:505–15, http://dx.doi.org/10.1016/j.cell.2007.08.046. [91] Branche C, Kohl L, Toutirais G, Buisson J, Cosson J, Bastin P. Conserved and specific functions of axoneme components in trypanosome motility. J. Cell Sci 2006;119:3443–55, http://dx.doi.org/10.1242/jcs.03078. [92] Baron DM, Kabututu ZP, Hill KL. Stuck in reverse: loss of LC1 in Trypanosoma brucei disrupts outer dynein arms and leads to reverse flagellar beat and backward movement. J. Cell Sci 2007;120:1513–20, http://dx.doi.org/10.1242/jcs.004846.

13

[93] Holwill MEJ. The motion of strigomonas oncopelti. J. Exp. Biol 1965;42:125–37. [94] Coles JA, Myburgh E, Ritchie R, Hamilton A, Rodgers J, Mottram JC, et al. Intravital imaging of a massive lymphocyte response in the cortical dura of mice after peripheral infection by trypanosomes. PLoS Negl. Trop. Dis 2015;9:e0003714, http://dx.doi.org/10.1371/journal.pntd.0003714. [95] Frevert U, Movila A, Nikolskaia OV, Raper J, Mackey ZB, Abdulla M, et al. Early invasion of brain parenchyma by African trypanosomes. PLoS ONE 2012;7:e43913, http://dx.doi.org/10.1371/journal.pone.0043913. [96] Matthews KR, McCulloch R, Morrison LJ. The within-host dynamics of African trypanosome infections. Philos. Trans. R. Soc. B 2015;370:20140288, http://dx.doi.org/10.1098/rstb.2014.0288. [97] Dean S, Marchetti R, Kirk K, Matthews KR. A surface transporter family conveys the trypanosome differentiation signal. Nature 2009;459:213–7, http://dx.doi.org/10.1038/nature07997. [98] Engstler M, Boshart M. Cold shock and regulation of surface protein trafficking convey sensitization to inducers of stage differentiation in Trypanosoma brucei. Genes Dev 2004;18:2798–811, http://dx.doi.org/10.1101/gad.323404. [99] Nolan DP, Rolin S, Rodriguez JR, Van Den Abbeele J, Pays E. Slender and stumpy bloodstream forms of Trypanosoma brucei display a differential response to extracellular acidic and proteolytic stress. Eur. J. Biochem 2000;267:18–27, http://dx.doi.org/10.1046/j.1432-1327.2000.00935.x. [100] Sharma R, Gluenz E, Peacock L, Gibson W, Gull K, Carrington M. The heart of darkness: growth and form of Trypanosoma brucei in the tsetse fly. Trends Parasitol 2009;25:517–24, http://dx.doi.org/10.1016/j.pt.2009.08.001. [101] Rotureau B, Ooi C-P, Huet D, Perrot S, Bastin P. Forward motility is essential for trypanosome infection in the tsetse fly. Cell. Microbiol 2014;16:425–33, http://dx.doi.org/10.1111/cmi.12230. [102] Dyer NA, Rose C, Ejeh NO, Acosta-Serrano A. Flying tryps: survival and maturation of trypanosomes in tsetse flies. Trends Parasitol 2013;29:188–96, http://dx.doi.org/10.1016/j.pt.2013.02.003. [103] Rotureau B, Subota I, Bastin P. Molecular bases of cytoskeleton plasticity during the Trypanosoma brucei parasite cycle. Cell. Microbiol 2011;13:705–16, http://dx.doi.org/10.1111/j.1462-5822.2010.01566.x. [104] Subota I, Rotureau B, Blisnick T, Ngwabyt S, Durand-Dubief M, Engstler M, et al. ALBA proteins are stage regulated during trypanosome development in the tsetse fly and participate in differentiation. Mol. Biol. Cell 2011;22:4205–19, http://dx.doi.org/10.1091/mbc.E11-06-0511. [105] Oberholzer M, Lopez MA, McLelland BT, Hill KL. Social motility in African trypanosomes. PLoS Pathog 2010;6:e1000739, http://dx.doi.org/10.1371/journal.ppat.1000739. [106] Imhof S, Knüsel S, Gunasekera K, Vu XL, Roditi I. Social motility of African trypanosomes is a property of a distinct life-cycle stage that occurs early in tsetse fly transmission. PLoS Pathog 2014;10:e1004493, http://dx.doi.org/10.1371/journal.ppat.1004493. [107] Alizadehrad D, Krüger T, Engstler M, Stark H. Simulating the complex cell design of Trypanosoma brucei and its motility. PLoS Comput. Biol 2015;11, http://dx.doi.org/10.1371/journal.pcbi.1003967. [108] Peacock L, Bailey M, Carrington M, Gibson W. Meiosis and haploid gametes in the pathogen Trypanosoma brucei. Curr. Biol 2014;24:181–6, http://dx.doi.org/10.1016/j.cub.2013.11.044. [109] Gibson W. Liaisons dangereuses: sexual recombination among pathogenic trypanosomes. Res. Microbiol 2015;166:459–66, http://dx.doi.org/10.1016/j.resmic.2015.05.005. [110] Portman N, Gull K. The paraflagellar rod of kinetoplastid parasites: from structure to components and function. Int. J. Parasitol 2010;40:135–48, http://dx.doi.org/10.1016/j.ijpara.2009.10.005. [111] Sunter JD, Varga V, Dean S, Gull K. A dynamic coordination of flagellum and cytoplasmic cytoskeleton assembly specifies cell morphogenesis in trypanosomes. J. Cell Sci 2015;128:1580–94, http://dx.doi.org/10.1242/jcs.166447. [112] Langousis G, Hill KL. Motility and more: the flagellum of Trypanosoma brucei. Nat. Rev. Microbiol 2014;12:505–18, http://dx.doi.org/10.1038/nrmicro3274. [113] Sinden R, Talman A, Marques S, Wass M, Sternberg M. The flagellum in malarial parasites. Curr. Opin. Microbiol 2010;13:491–500, http://dx.doi.org/10.1016/j.mib.2010.05.016. [114] Wilson LG, Carter LM, Reece SE. High-speed holographic microscopy of malaria parasites reveals ambidextrous flagellar waveforms. Proc. Natl. Acad. Sci 2013;110:18769–74, http://dx.doi.org/10.1073/pnas.1309934110. [115] Gray J, Hancock GJ. The propulsion of sea-urchin spermatozoa. J. Exp. Biol 1955;32:802–14. [116] Gray J. The movement of sea-urchin spermatozoa. J. Exp. Biol 1955;32:775–801. [117] Friedrich BM, Riedel-Kruse IH, Howard J, Julicher F. High-precision tracking of sperm swimming fine structure provides strong test of resistive force theory. J. Exp. Biol 2010;213:1226–34, http://dx.doi.org/10.1242/jeb.039800. [118] Lauga E, Powers TR. The hydrodynamics of swimming microorganisms. Rep. Prog. Phys 2009;72:096601, http://dx.doi.org/10.1088/0034-4885/72/9/096601.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15 14

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

[119] Purcell EM. Life at low Reynolds number. Am. J. Phys 1977;45:3, http://dx.doi.org/10.1119/1.10903. [120] Stellamanns E, Uppaluri S, Hochstetter A, Heddergott N, Engstler M, Pfohl T. Optical trapping reveals propulsion forces, power generation and motility efficiency of the unicellular parasites Trypanosoma brucei brucei. Sci Rep 2014;4, http://dx.doi.org/10.1038/srep06515. [121] Uppaluri S, Heddergott N, Stellamanns E, Herminghaus S, Zöttl A, Stark H, et al. Flow loading induces oscillatory trajectories in a bloodstream parasite. Biophys. J 2012;103:1162–9, http://dx.doi.org/10.1016/j.bpj.2012.08.020. [122] Kirkman-Brown JC, Smith DJ. Sperm motility: is viscosity fundamental to progress? Mol. Hum. Reprod 2011;17:539–44, http://dx.doi.org/10.1093/molehr/gar043. [123] Uppaluri S, Nagler J, Stellamanns E, Heddergott N, Herminghaus S, Engstler M, et al. Impact of microscopic motility on the swimming behavior of parasites: straighter trypanosomes are more directional. PLoS Comput. Biol 2011;7:e1002058, http://dx.doi.org/10.1371/journal.pcbi.1002058. [124] Berland K, Jacobson K, French T. Electronic cameras for low-light microscopy. Methods Cell Biol 1998;56:19–44. [125] O’Rourke B, Reibel DK, Thomas AP. High-speed digital imaging of cytosolic Ca2+ and contraction in single cardiomyocytes. Am. J. Physiol. – Heart Circ. Physiol 1990;259:H230–42. [126] Sanderson MJ. High-speed digital microscopy. Methods 2000;21:325–34, http://dx.doi.org/10.1006/meth.2000.1022. [127] Liu Z, Lavis LD, Betzig E. Imaging live-cell dynamics and structure at the single-molecule level. Mol. Cell 2015;58:644–59, http://dx.doi.org/10.1016/j.molcel.2015.02.033. [128] Cox S. Super-resolution imaging in live cells. Dev. Biol 2015;401:175–81, http://dx.doi.org/10.1016/j.ydbio.2014.11.025. [129] Fornasiero EF, Opazo F. Super-resolution imaging for cell biologists. BioEssays 2015;37:436–51, http://dx.doi.org/10.1002/bies.201400170. [130] Davies R, Graham J, Canepari M. Light sources and cameras for standard in vitro membrane potential and high-speed ion imaging. J. Microsc 2013;251:5–13, http://dx.doi.org/10.1111/jmi.12047. [131] Ross WN, Miyazaki K, Popovic MA, Zecevic D. Imaging with organic indicators and high-speed charge-coupled device cameras in neurons: some applications where these classic techniques have advantages. Neurophotonics 2015;2:021005, http://dx.doi.org/10.1117/1.NPh.2.2.021005. [132] Smith DR, Winters DG, Bartels RA. Submillisecond second harmonic holographic imaging of biological specimens in three dimensions. Proc. Natl. Acad. Sci. U. S. A 2013;110:18391–6, http://dx.doi.org/10.1073/pnas.1306856110. [133] Rodríguez JA, Lopez MA, Thayer MC, Zhao Y, Oberholzer M, Chang DD, et al. Propulsion of African trypanosomes is driven by bihelical waves with alternating chirality separated by kinks. Proc. Natl. Acad. Sci. U. S. A 2009;106:19322–7, http://dx.doi.org/10.1073/pnas.0907001106. [134] Garcia-Sucerquia J, Xu W, Jericho SK, Klages P, Jericho MH, Kreuzer HJ. Digital in-line holographic microscopy. Appl. Opt 2006;45:836–50, http://dx.doi.org/10.1364/AO.45.000836. [135] Sheng J, Malkiel E, Katz J. Digital holographic microscope for measuring three-dimensional particle distributions and motions. Appl. Opt 2006;45:3893–901, http://dx.doi.org/10.1364/AO.45.003893. [136] Cheong FC, Wong CC, Gao Y, Nai MH, Cui Y, Park S, et al. Rapid, high-throughput tracking of bacterial motility in 3D via phase-contrast holographic video microscopy. Biophys. J 2015;108:1248–56, http://dx.doi.org/10.1016/j.bpj.2015.01.018. [137] Di Caprio G, El Mallahi A, Ferraro P, Dale R, Coppola G, Dale B, et al. 4D tracking of clinical seminal samples for quantitative characterization of motility parameters. Biomed. Opt. Express 2014;5:690–700, http://dx.doi.org/10.1364/BOE.5.000690. [138] Weiße S, Heddergott N, Heydt M, Pflästerer D, Maier T, Haraszti T, et al. A quantitative 3D motility analysis of Trypanosoma brucei by use of digital in-line holographic microscopy. PLoS ONE 2012;7:e37296, http://dx.doi.org/10.1371/journal.pone.0037296. [139] van Duinen V, Trietsch SJ, Joore J, Vulto P, Hankemeier T. Microfluidic 3D cell culture: from tools to tissue models. Curr. Opin. Biotechnol 2015;35:118–26, http://dx.doi.org/10.1016/j.copbio.2015.05.002. [140] Esch EW, Bahinski A, Huh D. Organs-on-chips at the frontiers of drug discovery. Nat. Rev. Drug Discov 2015;14:248–60, http://dx.doi.org/10.1038/nrd4539. [141] Lee D-H, Bae CY, Kwon S, Park J-K. User-friendly 3D bioassays with cell-containing hydrogel modules: narrowing the gap between microfluidic bioassays and clinical end-users’ needs. Lab Chip 2015;15:2379–87, http://dx.doi.org/10.1039/C5LC00239G. [142] Knowlton SM, Sadasivam M, Tasoglu S. Microfluidics for sperm research. Trends Biotechnol 2015;33:221–9, http://dx.doi.org/10.1016/j.tibtech.2015.01.005. [143] Pushkarsky I, Liu Y, Weaver W, Su T-W, Mudanyali O, Ozcan A, et al. Automated single-cell motility analysis on a chip using lensfree microscopy. Sci. Rep 2014;4, http://dx.doi.org/10.1038/srep04717.

[144] Wu J, Zheng G, Lee LM. Optical imaging techniques in microfluidics and their applications. Lab Chip 2012;12:3566–75, http://dx.doi.org/10.1039/C2LC40517B. [145] Schuster FL, Sullivan JJ. Cultivation of clinically significant hemoflagellates. Clin. Microbiol. Rev 2002;15:374–89, http://dx.doi.org/10.1128/CMR.15.3.374-389.2002. [146] Duran-Rehbein GA, Vargas-Zambrano JC, Cuéllar A, Puerta CJ, Gonzalez JM. Mammalian cellular culture models of Trypanosoma cruzi infection: a review of the published literature. Parasite 2014;21:38, http://dx.doi.org/10.1051/parasite/2014040. [147] Cotton JA, Motta J-P, Schenck LP, Hirota SA, Beck PL, Buret AG. Giardia duodenalis infection reduces granulocyte infiltration in an in vivo model of bacterial toxin-induced colitis and attenuates inflammation in human intestinal tissue. PLoS ONE 2014;9:e109087, http://dx.doi.org/10.1371/journal.pone.0109087. [148] Duaso J, Rojo G, Cabrera G, Galanti N, Bosco C, Maya JD, et al. Trypanosoma cruzi induces tissue disorganization and destruction of chorionic villi in an ex vivo infection model of human placenta. Placenta 2010;31:705–11, http://dx.doi.org/10.1016/j.placenta.2010.05.007. [149] Antoine-Moussiaux N, Magez S, Desmecht D. Contributions of experimental mouse models to the understanding of African trypanosomiasis. Trends Parasitol 2008;24:411–8, http://dx.doi.org/10.1016/j.pt.2008.05.010. [150] Gomes-Silva A, Valverde JG, Ribeiro-Romão RP, Plácido-Pereira RM, Da-Cruz AM. Golden hamster (Mesocricetus auratus) as an experimental model for Leishmania (Viannia) braziliensis infection. Parasitology 2013;140:771–9, http://dx.doi.org/10.1017/S0031182012002156. [151] Adell AD, Miller WA, Harvey DJ, Van Wormer E, Wuertz S, Conrad PA. Individual subject meta-analysis of parameters for Giardia duodenalis shedding in animal experimental models. BioMed Res. Int 2014;2014:e476142, http://dx.doi.org/10.1155/2014/476142. [152] Cobo ER, Eckmann L, Corbeil LB. Murine models of vaginal trichomonad infections. Am. J. Trop. Med. Hyg 2011;85:667–73, http://dx.doi.org/10.4269/ajtmh.2011.11-0123. [153] Heussler V, Doerig C. In vivo imaging enters parasitology. Trends Parasitol 2006;22:192–5, http://dx.doi.org/10.1016/j.pt.2006.03.001. [154] Amino R, Thiberge S, Martin B, Celli S, Shorte S, Frischknecht F, et al. Quantitative imaging of Plasmodium transmission from mosquito to mammal. Nat. Med 2006;12:220–4, http://dx.doi.org/10.1038/nm1350. [155] Goyard S, Dutra PL, Deolindo P, Autheman D, D’Archivio S, Minoprio P. In vivo imaging of trypanosomes for a better assessment of host–parasite relationships and drug efficacy. Parasitol. Int 2014;63:260–8, http://dx.doi.org/10.1016/j.parint.2013.07.011. [156] Lewis MD, Fortes Francisco A, Taylor MC, Burrell-Saward H, McLatchie AP, Miles MA, et al. Bioluminescence imaging of chronic Trypanosoma cruzi infections reveals tissue-specific parasite dynamics and heart disease in the absence of locally persistent infection. Cell. Microbiol 2014;16:1285–300, http://dx.doi.org/10.1111/cmi.12297. [157] Taylor MC, Kelly JM. Optimizing bioluminescence imaging to study protozoan parasite infections. Trends Parasitol 2014;30:161–2, http://dx.doi.org/10.1016/j.pt.2014.01.006. [158] Thalhofer CJ, Graff JW, Love-Homan L, Hickerson SM, Craft N, Beverley SM, et al. In vivo imaging of transgenic leishmania parasites in a live host. J. Vis. Exp. JoVE 2010, http://dx.doi.org/10.3791/1980. [159] Van Reet N, Van de Vyver H, Pyana PP, Van der Linden AM, Büscher P. A panel of Trypanosoma brucei strains tagged with blue and red-shifted luciferases for bioluminescent imaging in murine infection models. PLoS Negl. Trop. Dis 2014;8:e3054, http://dx.doi.org/10.1371/journal.pntd.0003054. [160] Lang T, Goyard S, Lebastard M, Milon G. Bioluminescent Leishmania expressing luciferase for rapid and high throughput screening of drugs acting on amastigote-harbouring macrophages and for quantitative real-time monitoring of parasitism features in living mice. Cell. Microbiol 2005;7:383–92, http://dx.doi.org/10.1111/j.1462-5822.2004.00468.x. [161] Ustione A, Piston Dw. A simple introduction to multiphoton microscopy. J. Microsc 2011;243:221–6, http://dx.doi.org/10.1111/j.1365-2818.2011.03532.x. [162] Germain RN, Robey EA, Cahalan MD. A decade of imaging cellular motility and interaction dynamics in the immune system. Science 2012;336:1676–81, http://dx.doi.org/10.1126/science.1221063. [163] Coombes JL, Robey EA. Dynamic imaging of host–pathogen interactions in vivo. Nat. Rev. Immunol 2010;10:353–64, http://dx.doi.org/10.1038/nri2746. [164] Chong SZ, Evrard M, Ng LG. Lights, camera, and action: vertebrate skin sets the stage for immune cell interaction with arthropod-vectored pathogens. Microb. Immunol 2013;4:286, http://dx.doi.org/10.3389/fimmu.2013.00286. [165] Peters NC, Egen JG, Secundino N, Debrabant A, Kimblin N, Kamhawi S, et al. In vivo imaging reveals an essential role for neutrophils in Leishmaniasis transmitted by sand flies. Science 2008;321:970–4, http://dx.doi.org/10.1126/science.1159194.

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034

G Model YSCDB-1865; No. of Pages 15

ARTICLE IN PRESS T. Krüger, M. Engstler / Seminars in Cell & Developmental Biology xxx (2015) xxx–xxx

[166] Kolesnikov M, Farache J, Shakhar G. Intravital two-photon imaging of the gastrointestinal tract. J. Immunol. Methods 2015;421:73–80, http://dx.doi.org/10.1016/j.jim.2015.03.008. [167] Amino R, Thiberge S, Blazquez S, Baldacci P, Renaud O, Shorte S, et al. Imaging malaria sporozoites in the dermis of the mammalian host. Nat. Protoc 2007;2:1705–12, http://dx.doi.org/10.1038/nprot.2007.120. [168] Oreopoulos J, Berman R, Browne M. Spinning-disk confocal microscopy: present technology and future trends. Methods Cell Biol 2014;123:153–75, http://dx.doi.org/10.1016/B978-0-12-420138-5.00009-4. [169] Aurrecoechea C, Heiges M, Wang H, Wang Z, Fischer S, Rhodes P, et al. ApiDB: integrated resources for the apicomplexan bioinformatics resource center. Nucleic Acids Res 2007;35:D427–30, http://dx.doi.org/10.1093/nar/gkl880. [170] Meissner M, Agop-Nersesian Jr CWJS. Molecular tools for analysis of gene function in parasitic microorganisms. Appl. Microbiol. Biotechnol 2007;75:963–75, http://dx.doi.org/10.1007/s00253-007-0946-4. [171] Docampo R. Molecular parasitology in the 21st century. Essays Biochem 2011;51:1–13, http://dx.doi.org/10.1042/bse0510001. [172] Kolev NG, Tschudi C, Ullu E. RNA interference in protozoan parasites: achievements and challenges. Eukaryot. Cell 2011;10:1156–63, http://dx.doi.org/10.1128/EC.05114-11. [173] Dean S, Sunter J, Wheeler RJ, Hodkinson I, Gluenz E, Gull K. A toolkit enabling efficient, scalable and reproducible gene tagging in trypanosomatids. Open Biol 2015;5:140197, http://dx.doi.org/10.1098/rsob.140197.

15

[174] Peng D, Kurup SP, Yao PY, Minning TA, Tarleton RL. CRISPR-Cas9-mediated single-gene and gene family disruption in Trypanosoma cruzi. mBio 2015;6:e02097–2114, http://dx.doi.org/10.1128/mBio.02097-14. [175] Sollelis L, Ghorbal M, MacPherson CR, Martins RM, Kuk N, Crobu L, et al. First efficient CRISPR-Cas9-mediated genome editing in Leishmania parasites. Cell. Microbiol 2015;17:1405–12, http://dx.doi.org/10.1111/cmi.12456. [176] Lander N, Li Z-H, Niyogi S, Docampo R. CRISPR/Cas9-induced disruption of paraflagellar rod protein 1 and 2 genes in Trypanosoma cruzi reveals their role in flagellar attachment. mBio 2015;6:e01012–1015, http://dx.doi.org/10.1128/mBio.01012-15. [177] Santrich C, Moore L, Sherwin T, Bastin P, Brokaw C, Gull K, et al. A motility function for the paraflagellar rod of Leishmania parasites revealed by PFR-2 gene knockouts. Mol. Biochem. Parasitol 1997;90:95–109, http://dx.doi.org/10.1016/S0166-6851(97)00149-7. [178] Bastin P, Sherwin T, Gull K. Paraflagellar rod is vital for trypanosome motility. Nature 1998;391:548, http://dx.doi.org/10.1038/35300. [179] Dobell C, van Leeuwenhoek A. Antony van Leeuwenhoek and his “Little Animals”; Being Some Account of the Father of Protozoology and Bacteriology and his Multifarious Discoveries in These Disciplines. New York, Harcourt: Brace and Company; 1932. http://archive.org/details/ antonyvanleeuwen00dobe (accessed 11.08.15).

Please cite this article in press as: Krüger T, Engstler M. Flagellar motility in eukaryotic human parasites. Semin Cell Dev Biol (2015), http://dx.doi.org/10.1016/j.semcdb.2015.10.034