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Visualization of the funis of Giardia lamblia by high-resolution field emission scanning electron microscopy—new insights
Journal of
Structural Biology Journal of Structural Biology 147 (2004) 102–115 www.elsevier.com/locate/yjsbi
Visualization of the funis of Giardia lamblia by high-resolution field emission scanning electron microscopy—new insights Marlene Benchimol,a,* Bruno Piva,a Loraine Campanati,b and Wanderley de Souzab a
b
rsula, R. Jornalista Orlando Dantas, 59, Universidade Santa Ursula, Laboratorio de Ultraestrutura Celular—Universidade Santa U Rio de Janeiro, Brazil Universidade Federal do Rio de Janeiro, Instituto de Biofısica Carlos Chagas Filho, Laboratorio de Ultraestrutura Celular Hertha Meyer, CCS, Ilha do Fund~ao, 21940-900, Rio de Janeiro, Brazil Received 23 August 2003, and in revised form 5 December 2003 Available online 18 March 2004
Abstract Giardia lamblia is a multiflagellar parasite and one of the earliest diverging eukaryotic cells. It possesses a cytoskeleton made of several microtubular structures—an adhesive disc, four pairs of flagella, median body, and funis. This protozoan displays different types of movements, including a lateral and dorso-ventral dislocation of its posterior region, which has not been completely elucidated. In the present study, high-resolution field emission scanning electron microscopy was used to analyze the funis structure of G. lamblia trophozoites. It was shown that the funis is made of short arrays of microtubules emanating from the axonemes of the caudal flagella, which are anchored to dense rods that run parallel to the posterior-lateral flagella. After emergence of the posteriorlateral flagella, funis microtubules are anchored to the epiplasm, a fibrous layer that underlies the portion of membrane that presents tail contractility. Based on these observations a model for the tail flexion of G. lamblia is proposed. Ó 2004 Elsevier Inc. All rights reserved. Keywords: Giardia lamblia; Funis; FESEM; Cell motility; Microtubules
1. Introduction Giardia lamblia is a parasitic protozoan that infects thousands of people all over the world, causing a disease known as giardiasis. The trophozoite form of this protist lacks organelles usually found in higher eukaryotes, such as mitochondria and peroxisomes (Gillin et al., 1996). From studies on rRNA structure of G. lamblia, which has been found to be close to that of archaebacteria, this organism has been considered to be a suitable model for the study of evolution of cell structures (Edlind and Chakraborty, 1987; Sogin et al., 1989). It presents an unique cytoskeleton in which the protein tubulin predominates, specially in the following structures: four pairs of flagella, an adhesive disc composed of microtubules and microribbons containing giardins, a median body, and a funis made up of sheets of micro* Corresponding author. Fax: +55-21-2553-1615. E-mail address: [email protected] (M. Benchimol).
1047-8477/$ - see front matter Ó 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.jsb.2004.01.017
tubules following the axonemes of the caudal flagella (Campanati et al., 2002; Erlandsen and Feely, 1984; Kulda and Nohynkova, 1995; Upcroft and Upcroft, 1998). Among these structures the funis and the median body are the least defined and deserve investigation, since their function have not been yet clarified. Giardia has eight flagella, arranged in four pairs termed anterior, posterior-lateral, ventral, and caudal. Both the anterior and posterior-lateral flagella are followed on their posterior side by electron dense material, termed dense or fibrous rods (Erlandsen and Feely, 1984; Kulda and Nohynkova, 1995). Giardia motility has been recently studied by videomicroscopy (Campanati et al., 2002; Ghosh et al., 2001). It was shown that the anterior and ventral flagella present characteristic patterns and frequencies of beating, while the posterior and the caudal one do not show a wave-beating pattern. As documented by microcinematography (Erlandsen and Feely, 1984), and videomicroscopy (Campanati et al., 2002; Ghosh et al., 2001),
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the tail displays a lateral rocking movement and dorso-ventral flexion. Campanati et al. (2002) showed that motility of the caudal region was completely independent from the flagellar beating and it was likely to be produced by a microtubular complex located in the caudal portion of the cell. The funis is a poorly studied microtubular structure. It was described by Holberton (1973) as short arrays of microtubules emanating from the axonemes of the caudal flagella, one in the dorsal and one in the ventral position, and was designated as funis by Kulda and Noh ynkov a (1978). In the internuclei area the funis consists of well-defined bands of interconnected microtubules and from the point of emergence of the ventral flagella, the individual microtubules detach from each band and fan out laterally in a position dorsal to the posterior-lateral axonemes, down to the tail of the organism (Campanati et al., 2003; Kulda and Noh ynkova, 1978). It has been suggested that the funis would have a structural function (Kulda and Noh ynkov a, 1995). The aim of this paper is to describe the fine structure of the funis, its connections, and its possible functional role. High-resolution scanning electron microscopy images of the funis showed that its microtubules do not end in the cytoplasm, but are anchored in the posterior flagella, via the dense rods. In addition, after the emergence of the posterior-lateral flagella, the funis microtubules irradiate towards the plasma membrane and attach to filamentous links underlying the membrane. We suggest that the funis participates on the lateral tail flexion and propose a model for the caudal motility of G. lamblia.
2. Materials and methods 2.1. Organisms and culture Giardia lamblia WB strain (American Type Culture Collection, No. 30957) was cultivated in TYI-S-33 medium enriched with 10% heat-inactivated bovine serum (Diamond et al., 1978) at pH 7.05, without added vitamins, iron, or antibiotics (Gillin et al., 1989), but supplemented with 0.1% bovine bile (Keister, 1983) for 48–72 h, at 37 °C.
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OsO4 in cacodylate buffer containing 5 mM CaCl2 and 0.8% potassium ferricyanide in the dark, for 30 min. Cells were washed in PBS, dehydrated in acetone, and embedded in Epon. Ultra-thin sections were stained with uranyl acetate for 20 min and lead citrate for 5 min and observed in a JEOL 1210 or a Zeiss 900 transmission electron microscope operating at 80 kV. All reagents, unless otherwise stated, were purchased from Sigma, St. Louis, USA. For negative staining PBS washed cells were allowed to adhere to formvar coated grids for 2 min, and the membranes were extracted in a solution containing 1% Nonidet P-40 in PHEM buffer (Schliwa and van Blerkom, 1981) for 40 min. After washes in PHEM buffer, the samples were fixed in 2% glutaraldehyde, washed with water, and then stained with 1% ammonium molibdate for 1 min. Next, the grids were observed in a Zeiss 900 TEM. 2.3. Conventional scanning electron microscopy Living cells were adhered to poly-L -lysine-coated glass coverslips, fixed with 2.5% glutaraldehyde in cacodylate buffer, post-fixed for 5 min in 1% OsO4 , dehydrated in ethanol, critical point dried with CO2 , sputter-coated with gold–palladium, and examined in a Jeol 5800 scanning electron microscope operating at 12 kV. 2.4. High-resolution field emission scanning electron microscopy Cells were adhered to poly-L -lysine coated glass coverslips and then treated with the permeabilization buffer (0.5% Nonidet P-40, 0.1 M Pipes, 1 mM MgSO4 , 2 mM glycerol, 2 mM EGTA, 1 mM PMSF (phenylmethylsulfonyl fluoride), and 0.5% Triton X-100) for different times (10 min–2 h). The cells were washed in PBS and then fixed in 2.5% glutaraldehyde in phosphate buffer, post-fixed for 5 min in 1% OsO4 , dehydrated in ethanol, critical point dried with CO2 , and sputtercoated with carbon. The samples were examined in a Jeol JSM-6340F Field Emission Scanning Electron Microscope operated at an accelerating voltage of 5 kV, using 5 mm as working distance and the standard SEI and BEI detectors.
2.2. Transmission electron microscopy1 2.5. Immunoelectron microscopy Cells were fixed overnight at room temperature in 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2). Then, the cells were washed twice in PBS (phosphatebuffered saline). Post-fixation was performed in 1% 1
Abbreviations used: FESEM, field emission scanning electron microscopy; SEM, scanning electron microscopy; TEM, transmission electron microscopy.
Cells were adhered to poly-L -lysine coated glass coverslips and then treated with the permeabilization buffer as above described. For immunolabeling, the cytoskeletons were first incubated in a solution of PBS containing 3% albumin, quenched in 50 mM NH4 Cl for 30 min, and incubated overnight in the presence of a monoclonal anti-a-tubulin TAT-1 antibody, kindly
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provided by Dr. K. Gull (Woods et al., 1989). After several washes in PBS–1% albumin, the cytoskeletons were incubated in the presence of 15 nm gold-labeled goat anti-mouse IgG (BBInternational, UK). Coverslips were washed several times in PBS and then fixed in 2.5% glutaraldehyde in phosphate buffer, post-fixed for 15 min in 1% OsO4 , dehydrated in ethanol, critical point dried with CO2 , and sputter-coated with carbon. The samples were examined in a Jeol JSM-6340F Field
Emission Scanning Electron Microscope operated at an accelerating voltage of 5 kV, using 5 mm as working distance, and the standard SEI and BEI detectors.
3. Results When observed by routine transmission or scanning electron microscopy Giardia trophozoites showed a
Fig. 1. Routine preparation for transmission electron microscopy of G. lamblia showing the ventral disc (D), the two nuclei (N), peripheral vesicles (V), flagellar axonemes (A), and the funis (arrows). (a) Shows a cross-section where the funis is seen as linear microtubular structures following the caudal flagella (arrows). (b,c) Longitudinal sections showing the funis microtubules (Fn) emanating from the caudal flagella (C). In (c) the brigdes between the funis microtubules are pointed by arrow (inset). MB, median body. Bars ¼ 1 lm.
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half-pear or teardrop shaped body (Figs. 1b, 3a). The ventral disc was seen in the anterior region of the cell and it was laterally and anteriorly surrounded by the marginal groove and the ventrolateral flange (Figs. 1a and 2). Gi-
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ardia presented four pairs of flagella, namely the anterior, posterior-lateral, caudal, and ventral (Figs. 2 and 3a,b). Ultrastructural examination of detergent-treated G. lamblia by FESEM allowed the visualization of its
Fig. 2. Negative staining of G. lamblia using 1% ammonium molibdate. Giardia was extracted by detergents (see Section 2), allowed to adhere on a formvar-coated grid, stained, and observed in the transmission electron microscope. The funis (arrow) is seen, as well the disc (D), and flagella. The nuclei (N) are dorsally situated in relation to the disc. Bar ¼ 1 lm.
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Fig. 3. Routine scanning electron microscopy (a) and field emission scanning electron microscopy (FESEM) after extraction with Triton X-100 (see Section 2) (b–d). In (b) the funis (arrow) is observed as microtubules connecting the axonemes of the caudal flagella (C) to the posterior-lateral flagella (P). The adhesive disc (D) and the four pairs of flagella, the ventral (V), posterior-lateral (P), anterior (A), and caudal (C) are seen. In (c) the caudal region shows the funis microtubules (Fn) emanating from the caudal flagella (C) to the posterior-lateral flagella (P). (c) A higher magnification of the caudal region seen in (b). Note that the funis microtubules originating from the right caudal flagellum cover the left caudal flagellum before reaching the fibrillar material. After the emergence of the posterior-lateral flagella the microtubules are directly connected to the fibrillar material underlying the plasma membrane. In (d), a close view of the caudal flagella (C) from where the funis microtubules (Fn) fan out towards the posteriorlateral flagella (P) and are seen anchored to the dense rods (arrows). Note some of the filamentous bridges interconnecting the funis microtubules. Bars ¼ 1 lm.
microtubular cytoskeleton (Fig. 2). The adhesive disc, located in the ventral anterior region of the trophozoite, was mainly made of concentrically arranged microtubules (Figs. 1a, 2, 3a, 4a, and 5). The disc microtubules and their associated dorsal ribbons were organized in a spiralized organization, surrounding a central area devoid of microtubules, known as the bare area (Figs. 1a, 3a, and 4a). The thickness of the microtubules observed in our preparations varied as a function of the carbon layer thickness used in the sample preparation and also by a coat of fibrous material present in some microtubular structures. This can be compared observing the microtubules of the funis, and the median body (Fig. 6b).
The axonemes of the caudal flagella present emanating microtubules (Figs. 1b,c, 2, 3b–d, and 4–8), which are part of a structure named funis by Kulda and Nohynkova (1978). At the most anterior region of the caudal flagella, near the basal bodies, the microtubules of the funis wrapped the caudal axonemes (Figs. 3d and 8a). Gradually, these microtubules begin to leave the caudal axonemes and fan out towards the axonemes of the posterior-lateral flagella (Figs. 3b–d, 4a,b, and 5–7). These microtubules were interconnected by bridges and attached to the fibrous rod (dense rod) present on the axoneme of the posterior-lateral flagella (Figs. 3d and 6b). Thus, the caudal and posterior-lateral axonemes flagella become linked by the funis microtubules (Figs.
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Fig. 4. High-resolution field emission scanning electron microscopy of Giardia cytoskeleton showing microtubules emanating from the caudal flagella towards the plasma membrane (arrow). Adhesive disc (D); funis (Fn); anterior flagella (A); posterior-lateral flagella (P); ventral flagella (V); and caudal flagella (C). (b) A close view of the ventral region showing the funis microtubules (Fn) which are linked by filamentous bridges (arrow). Posterior-lateral flagella (P); caudal flagella (C); ventral flagella (V); and ventral disc (D). Bars ¼ 1 lm.
3b,c, 5, and 6). When Giardia was adhered through its ventral region (adhesive disc down) on poly-L -lysine coated coverslips, the funis was observed on its dorsal face (Fig. 5). This view allowed the observation that the funis microtubules arised from one of the caudal flagella axonemes, running towards the left, contacting the other caudal flagellum axoneme and adhering to the dense rods of the posterior-lateral flagellum axoneme (Fig. 5). When Giardia was adhered through its dorsal region, (adhesive disc up) it was possible to observe that the other caudal flagellum axoneme also presented emanating microtubules, which are directed to the opposite posterior-lateral flagellar axoneme (Figs. 3, 4, and 6). Thus, there are two emanating groups of microtubules from the axonemes of the caudal flagella that migrate in opposite directions, forming a well-structured array (Figs. 3–6, and 10). In addition, the funis microtubules were covered by an unidentified material, which gave them a greater thickness. When cytoskeletons were isolated by suspending cells in Triton without first attaching them to a surface, several structures, such as microtubules, bridges, and the median bodies were normally lost into the supernatant. Crossley et al. (1986) have made a similar observation. Thus, several trials were made using different times and
detergent concentrations until a good preservation of the bridges was achieved. Only in cells well extracted by detergent the funis were clearly seen. The microtubules of the funis presented bridges (Figs. 1c, 3d, and 6b), which were very sensitive to Triton X-100, and were easily lost during cytoskeleton preparations. By means of the funis microtubules the caudal flagella axonemes seems to be firmly anchored to the axoneme of the posterior-lateral flagella. In addition, after the emergence of the posterior-lateral flagella, few microtubules were seen projecting towards the plasma membrane. They were anchored to a network found underlying the membrane (Fig. 3d). These structures were preserved during sample preparation, using different times of extraction and detergent concentrations, although the bridges interconnecting the funis microtubules were affected when longer times, high detergent concentrations, or centrifugation were used (Fig. 3). The microtubules of the median body were dorsally positioned in relation to the funis and were seen as smooth cylinders, rather distinct from those of the funis, which are covered by a rough coat (Fig. 6). A linearly aligned material, forming images similar to ‘‘corn kernels,’’ covered the posterior-lateral flagella. This coating material was rough and the grains were
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Fig. 5. Scanning electron microscopy showing a dorsal view of the funis microtubules (Fn). They emanate from each of the caudal flagellum (C), contact the other flagellum (arrow) and attach to the dense rod of the opposite posterior-lateral flagellum (P). D, disc; A, anterior flagellum. Note that the arrow and the arrowhead point to funis microtubules found in distinct levels. Bar ¼ 1 lm.
seen in linear rows (Fig. 7). Immunocytochemistry was performed after the detergent extraction and monoclonal anti-tubulin TAT-1, was used (Fig. 8). The funis
microtubules were intensely labeled and were better visualized when back-scattered electrons were used (BEI), as shown in Fig. 8b.
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Fig. 6. (a) SEM of G. lamblia in a ventral view. Note that the microtubules of the funis (Fn) emanate from the caudal flagella (C) and are anchored to the posterior-lateral flagella (P). The median body (MB) is also observed as bundles of microtubules close to the funis. The arrows point to nuclei prints which are located dorsally to the ventral adhesive disc (D). Bar ¼ 1 lm. (b) FESEM of G. lamblia in a close view. The funis microtubules (Fn) are observed emanating from the caudal flagella (C) towards the posterior-lateral flagella (P). Notice that the microtubules present links (arrows), many of which were disrupted by the extraction treatment. The median body (MB), also formed by microtubules, is seen in a dorsal position in relation to the funis. Ventral flagella (V). Bar ¼ 100 nm.
After the emergence of the posterior-lateral flagella, the funis microtubules contacted the plasma membrane (Figs. 3b,c and 4a), via a fibrilar material that underlies the membrane. It is well-known that Giardia presents a tail movement (Fig. 9). When cells are well-fixed the caudal region can be seen in a bent position, either by routine SEM (Fig. 9a) or after detergent extraction (Fig. 9b). We propose that the funis microtubules play some role on this type of movement, as suggested in a schematic drawing (Fig. 10).
4. Discussion Giardia has a complex cytoskeleton formed by different groups of structures that participate in important cell functions, such as adhesion and motility. Although many researchers have investigated motility and attachment processes in Giardia, findings concerning the role of the cytoskeleton in several processes are scant or contradictory. In addition, two components of the in Giardia cytoskeleton received almost no attention: the funis and the median body. Herein we show by the first time images of the cytoskeleton of G. lamblia as visualized by scanning electron
microscopy, allowing to get new insights into several structures. We made use of high-resolution field emission scanning electron microscopy (FESEM) and conventional SEM associated to plasma membrane extraction using different detergent concentrations, and immunogold labeling in order to get #new views of Giardia ultrastructure. Different times, types and detergents concentrations, as well several details on the centrifugation steps were tried before to achieve these preparations. The caudal microtubular structure termed funis (Kulda and Nohynkova, 1978) was carefully analyzed. The funis was described in the literature as a girdle of parallel single tubules that partly enclose the caudal axonemes (Crossley et al., 1986), and as sheets of microtubules (Campanati et al., 2002; Kulda and Nohynkova, 1995). It was considered that the funis microtubules provided a set of lateral ribs reaching the posterior cytoplasm (Campanati et al., 2003; Crossley et al., 1986; Kulda and Nohynkova, 1995). Here we confirmed that the funis is in fact made of microtubules, since they were labeled with anti-tubulin monoclonal antibodies. In addition, we show by the first time the connections between the funis microtubules and the posterior-lateral flagella, suggesting this participation on
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Fig. 7. High-resolution field emission scanning electron microscopy of G. lamblia in high magnification. The funis microtubules (Fn) are clearly seen anchored to the posterior-lateral flagella (P). Adhesive disc (D); ventral flagella (V). Bar ¼ 1 lm.
the Giardia caudal movement. All previous studies proposed that the funis did not present any structural connection when they emerge from the caudal flagella (Crossley et al., 1986; Kulda and Noh ynkov a, 1978, 1995). FESEM observations of the funis showed that this structure is formed by several single interconnected microtubules linking the caudal to the posterior-lateral flagella. Lateral microtubules emanate from each caudal flagellum, contact the other caudal flagellum and attach to the dense rod of the opposite posterior-lateral flagellum. The dorsal group of microtubules arises from the left caudal axoneme, close to the basal body, crosses to the right side of the cell, whereas the ventral group arises on the right, and reaches the left side. Each group of the funis microtubules runs in opposite directions and thus seems to exert more resistance to stress conditions. After the emergence of the posteriorlateral flagella the funis microtubules are seen anchored to the underlying material in the plasma membrane. Many protists posses a unique cortical cytoplasm: the epiplasm, which is a prominent proteinaceous layer
underlying the plasma membrane (Huttenlauch and Stick, 2003). Thus, the funis microtubules seem to be anchored to the epiplasm in the most posterior region of Giardia. It is this region that presents a lateral tail movement. By using our new protocol, new information were obtained on Giardia cytoskeleton. The funis microtubules are interconnected by bridges and coated by an undefined material. On the other hand, one of the microtubules ends is not freely dispersed in the cytoplasm, as proposed in previous studies, but firmly connected to posterior-lateral flagella or to the plasma membrane, after the emergence of the flagella. There are numerous examples in the literature of bridges connecting microtubules (McIntosch, 1974), which are important for maintaining the order of the microtubule array. In several cases such association is involved in the generation of motility. Motor proteins, which power cytoskeletal movements, have been found in other organisms. However, microtubule-based motor proteins, such as kinesin and dynein, and the microfilament-based myosin, are virtually uncharacterized in
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Fig. 8. High-resolution field emission scanning electron microscopy of G. lamblia after immunogold labeling using the monoclonal antibody TAT-1. In (a), the image is seen by SEI (a) whereas in (b) the same cell is shown by BEI. It is possible to see the gold particles labeling the microtubules of the adhesive disc (D), funis (Fn), caudal (C), and posterior-lateral flagella (P). Bar ¼ 1 lm.
Giardia (Elmendorf et al., 2003). Campanati et al. (2002) using high-pressure freezing and freeze-substitution observed arms emanating from the funis micro-
tubules. They assumed that the tiny filaments attached to the microtubules of these sheets might be dynein molecules. However, immunocytochemistry location of
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Fig. 9. Conventional scanning electron microscopy of G. lamblia seen on dorsal view, without membrane extraction (a) and in ventral view, after membrane extraction by detergents (b). Notice the caudal bent. D, disc; P, posterior-lateral flagella; C, caudal flagella; A, anterior flagella; V, ventral flagella; and Fn, funis. Bars ¼ 1 lm.
this protein using commercial antibodies was unsuccessful. In the present report, we observed these arms more clearly, and they were also contacting the neighbor microtubules.
The parasite motion is complex. Giardia displays different types of movements either when swimming or attached. The four pairs of flagella control different aspects of motility (Elmendorf et al., 2003). While the
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Fig. 10. Schematic drawing of a static G. lamblia (a,b) and presenting tail movement (c) showing the relationship of the funis and the posterior-lateral and caudal flagella. The caudal flagella pair (C1,C2) present an array of microtubules (in lilac and green) that emanate towards the posterior-lateral flagella (P1,P2). Note that in the caudal flagellum C2 the microtubules emanate towards the dense rods of the posterior-lateral flagellum 1 (P1), whereas the caudal flagellum 1 (C1) the microtubules are directed towards the posterior flagellum 2 (P2). Notice that the funis microtubules are anchored to the dense rods (in red) of the posterior-lateral flagella, and are interconnected by filamentous bridges (orange). After the emergence of the posterior-lateral flagella the funis microtubules are anchored to the epiplasm, a layer of material underlying the plasma membrane. This region presents a network of filamentous structures (F). The pair of caudal flagella is interconnected by thin bridges. During tail flexion (c) the microtubules of the funis found after the posterior-lateral flagella emergence are seen anchored to the material underlying the plasma membrane and presenting different length. They seem to participate in the caudal movement. D, disc; PV, peripheral vesicles. Bars ¼ 1 lm.
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parasite is attached, the ventral flagella beat continuously and when the cells are freely moving they present either a lateral rocking or a complete rotational (tumbling) movement along its longitudinal axis (Campanati et al., 2002; Elmendorf et al., 2003). An interesting phenomenon observed in G. lamblia is the movement performed by its tail region. This portion of the cell is able to bend both laterally and/or dorsoventrally. This movement is completely independent from the flagellar beating (Campanati et al., 2002; Glesbki, 1967). It was proposed that the microtubular complex located in the caudal portion of the cell likely produced the dislocation of the caudal region (Campanati et al., 2002). However, even performing highpressure freezing of living cells and freeze-substitution, structural details of this caudal region were not achieved in previous studies (Campanati et al., 2002). In the present study, FESEM associated with a detergent extraction protocol, allowed a better understanding of this cell region. The mechanism of dorsal and lateral flexion is not completely understood. It was suggested that this flexion is structurally associated in some way with the caudal flagella (Campanati et al., 2002), but it remains to be determined whether this force is derived only from flagellar movement or also from contractile proteins, associated in some fashion with the flagellar axonemes or the funis (Erlandsen and Feely, 1984). The structural organization of funis microtubules observed in this study perfectly fits with the observations made by Campanati et al. (2002) using video-microscopy in which the caudal flagella axonemes were seen rolling over each other, during the lateral dislocation of the caudal region of the cell. Dense fibrillar material has been reported by Feely et al. (1982) to parallel the course of the caudal flagella and their accompanying microtubules (funis), but the nature of this fibrillar material is unknown. Feely et al. (1982) found actin and a-actinin staining by immunofluorescence microscopy in the region corresponding to the dense rod-like masses near the axonemes of the posterior-lateral flagella and Narcisi et al. (1994) found actin along the intracellular axonemes. Feely et al. (1982) raised the possibility that the association of contractile proteins and microtubules could have functional significance. Careful observation of their images suggested us an intense labeling in the area of the funis. This finding, if confirmed, could explain the caudal contractility. Contractile proteins could participate, together with the funis microtubules, in the tail flexion. We observed that both the caudal and posterior-lateral flagella present a coat made of an uncharacterized material. The posterior-lateral flagella presented this dense coat covering their axonemes, whereas in the caudal flagella the coat covers only some areas, probably the regions of higher stress. In addition, a dense material
also covers the caudal flagella. It is necessary to determine the nature of the proteins composing these dense rods. Campanati et al. (2002) observed by video-microscopy that during the lateral movement of the caudal region one axoneme seemed to roll over the other one. They proposed that the movement generated by the inner axonemal apparatus could be constrained by the accessory microtubular structures and by the other axoneme, producing the movement of the caudal region. Our observations showed that indeed one group of microtubules originated from one of the caudal flagellum running over the other flagellum and in this course established contact with the other caudal flagellum (see scheme in Fig. 10). Like the interaction of microtubules in the axoneme, a sliding of one microtubule over the other could lead to shortening of the funis and, as consequence, the tail movement. We propose that the funis works as a stress and flexible cord. Thus, the name funis proposed by Kulda and Nohynkova (1978) is adequate (it means rope, cord). We speculate that the funis could be responsible for the caudal lateral movement in G. lamblia. This could be accomplished by the sliding of a microtubule over the other, as occurs in movements of flagellar axonemes. In addition, contractile proteins associated with funis microtubules could also participate in this movement. Another hypothesis includes a rapid polymerization/ depolymerization of microtubules in the region of insertion to the dense rods in the posterior-lateral flagellum, leading to caudal lateral flexion in Giardia.
Acknowledgments This work was supported by the Conselho Nacional de Desenvolvimento Cientıfico e Tecnol ogico (CNPq), Fundacß~ao Carlos Chagas Filho de Amparo a Pesquisa do Estado do Rio de Janeiro (FAPERJ), Programa de N ucleos de Excel^encia (PRONEX), Coordenacß~ ao de Aperfeicßoamento de Pessoal de Nıvel Superior (CAPES), and Associacß~ao Universitaria Santa Ursula (AUSU). The authors thank the technical support of William Christian Mol^edo Lopes.
References Campanati, L., Holloschi, A., Troster, H., Spring, H., De Souza, W., Monteiro-Leal, L.H., 2002. Video-microscopy observations of fast dynamic processes in the protozoon Giardia lamblia. Cell Motil. Cytoskel. 51, 213–224. Campanati, L., Troster, H., Monteiro-Leal, L.H., Spring, H., Trendelenburg, M.F., De Souza, W., 2003. Tubulin diversity in trophozoites of Giardia lamblia. Histochem. Cell Biol. 119, 323–331. Crossley, R., Marshall, J., Clark, J.T., Hooberton, D.V., 1986. Immunocytochemical differentiation of microtubules in the cytoskeleton of Giardia lamblia using monoclonal antibodies to
M. Benchimol et al. / Journal of Structural Biology 147 (2004) 102–115 a-tubulin and polyclonal antibodies to associated low molecular weight proteins. J. Cell Sci. 80, 233–252. Diamond, L.S., Harlow, D., Cunnick, C.C., 1978. A new medium for the axenic cultivation of Entamoeba histolytica and other Entamoeba. Trans. R. Soc. Trop. Med. Hyg. 72, 431–432. Edlind, T.D., Chakraborty, P.R., 1987. Unusual ribosomal RNA of the intestinal parasite Giardia lamblia. Nucleic Acids Res. 15, 7889– 7901. Elmendorf, G.H., Dawson, S.C., McCaffery, J.M., 2003. The cytoskeleton of Giardia lamblia. Int. J. Parasitol. 33, 3–28. Erlandsen, S.L., Feely, D.E., 1984. Trophozoite motility and the mechanism of attachment. In: Erlandsen, S.L., Meyer, E.A. (Eds.), Giardia and Giardiasis. Plenum, New York and London, pp. 33–63. Feely, D.E., Schollmeyer, J.V., Erlandsen, S.L., 1982. Giardia spp.: distribution of contractile proteins in the attachment organelle. Exp. Parasitol. 53, 145–154. Gillin, F.D., Reiner, D.S., McCaffery, J.M., 1996. Cell biology of the primitive eukaryote Giardia lamblia. Ann. Rev. Microbiol. 50, 679–705. Gillin, F.D., Boucher, S.E., Rossi, S.S., Reiner, D.S., 1989. Giardia lamblia the roles of bile, lactic acid and pH in completion of the life cycle in vitro. Exp. Parasitol. 69, 164–174. Ghosh, S., Frisardi, M., Rogers, R., Samuelson, J., 2001. How Giardia swim and divide. Infect. Immun. 69, 7866–7872. Glesbki, J., 1967. The movement in the cell Lamblia intestinalis. Acta Parasitol. Pol. 15, 109–111. Holberton, D.V., 1973. Fine structure of the ventral disc apparatus and the mechanism of attachment in the flagellate Giardia muris. J. Cell Sci. 13, 11–41.
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Huttenlauch, I., Stick, R., 2003. Occurrence of articulins and epiplasmins in protists. J. Euk. Microbiol. 50, 15–18. Keister, D.B., 1983. Axenic cultivation of Giardia lamblia in TYI-S-33 medium supplemented with bile. Trans. R. Soc. Trop. Med. Hyg. 77, 487–488. Kulda, J., Nohynkova, E., 1978. Flagellates of the human intestine and of intestines of other species. In: Kreier, J.P. (Ed.), Parasitic Protozoa. Intestinal Flagellates: Histomonads, Trichomonads, Amoeba, Opalinids, and Ciliates, vol. II. Academic Press, New York, pp. 2–139. Kulda, J., Nohynkova, E., 1995. Giardia in humans and animals, second ed.. In: Kreier, J.P. (Ed.),. Parasitic Protozoa, vol. 10. Academic Press, San Diego, pp. 225–422. McIntosch, J.R., 1974. Bridges between microtubules. J. Cell Biol. 61, 166–187. Narcisi, E.M., Paulin, J.J., Fechheimer, M., 1994. Presence and localization of vinculin in Giardia. J. Parasitol. 80, 468–473. Schliwa, M., van Blerkom, J., 1981. Structural interaction of cytoskeletal components. J. Cell Biol. 90, 222–235. Sogin, M.L., Gunderson, J.H., Elwood, H.J., Alonso, R.A., Peattie, D.A, 1989. Phylogenetic meaning of the kingdom concept: an usual ribosomal RNA from Giardia lamblia. Science 243, 75–77. Upcroft, J., Upcroft, P., 1998. My favorite cell: Giardia. BioEssays 20, 256–263. Woods, A., Sherwin, T., Sasse, R., MacRae, T.H., Baines, A.J., Gull, K., 1989. Definition of individual components within the cytoskeleton of Trypanosoma brucei by a library of monoclonal antibodies. J. Cell Sci. 93, 491–500.
Structural Biology Journal of Structural Biology 147 (2004) 102–115 www.elsevier.com/locate/yjsbi
Visualization of the funis of Giardia lamblia by high-resolution field emission scanning electron microscopy—new insights Marlene Benchimol,a,* Bruno Piva,a Loraine Campanati,b and Wanderley de Souzab a
b
rsula, R. Jornalista Orlando Dantas, 59, Universidade Santa Ursula, Laboratorio de Ultraestrutura Celular—Universidade Santa U Rio de Janeiro, Brazil Universidade Federal do Rio de Janeiro, Instituto de Biofısica Carlos Chagas Filho, Laboratorio de Ultraestrutura Celular Hertha Meyer, CCS, Ilha do Fund~ao, 21940-900, Rio de Janeiro, Brazil Received 23 August 2003, and in revised form 5 December 2003 Available online 18 March 2004
Abstract Giardia lamblia is a multiflagellar parasite and one of the earliest diverging eukaryotic cells. It possesses a cytoskeleton made of several microtubular structures—an adhesive disc, four pairs of flagella, median body, and funis. This protozoan displays different types of movements, including a lateral and dorso-ventral dislocation of its posterior region, which has not been completely elucidated. In the present study, high-resolution field emission scanning electron microscopy was used to analyze the funis structure of G. lamblia trophozoites. It was shown that the funis is made of short arrays of microtubules emanating from the axonemes of the caudal flagella, which are anchored to dense rods that run parallel to the posterior-lateral flagella. After emergence of the posteriorlateral flagella, funis microtubules are anchored to the epiplasm, a fibrous layer that underlies the portion of membrane that presents tail contractility. Based on these observations a model for the tail flexion of G. lamblia is proposed. Ó 2004 Elsevier Inc. All rights reserved. Keywords: Giardia lamblia; Funis; FESEM; Cell motility; Microtubules
1. Introduction Giardia lamblia is a parasitic protozoan that infects thousands of people all over the world, causing a disease known as giardiasis. The trophozoite form of this protist lacks organelles usually found in higher eukaryotes, such as mitochondria and peroxisomes (Gillin et al., 1996). From studies on rRNA structure of G. lamblia, which has been found to be close to that of archaebacteria, this organism has been considered to be a suitable model for the study of evolution of cell structures (Edlind and Chakraborty, 1987; Sogin et al., 1989). It presents an unique cytoskeleton in which the protein tubulin predominates, specially in the following structures: four pairs of flagella, an adhesive disc composed of microtubules and microribbons containing giardins, a median body, and a funis made up of sheets of micro* Corresponding author. Fax: +55-21-2553-1615. E-mail address: [email protected] (M. Benchimol).
1047-8477/$ - see front matter Ó 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.jsb.2004.01.017
tubules following the axonemes of the caudal flagella (Campanati et al., 2002; Erlandsen and Feely, 1984; Kulda and Nohynkova, 1995; Upcroft and Upcroft, 1998). Among these structures the funis and the median body are the least defined and deserve investigation, since their function have not been yet clarified. Giardia has eight flagella, arranged in four pairs termed anterior, posterior-lateral, ventral, and caudal. Both the anterior and posterior-lateral flagella are followed on their posterior side by electron dense material, termed dense or fibrous rods (Erlandsen and Feely, 1984; Kulda and Nohynkova, 1995). Giardia motility has been recently studied by videomicroscopy (Campanati et al., 2002; Ghosh et al., 2001). It was shown that the anterior and ventral flagella present characteristic patterns and frequencies of beating, while the posterior and the caudal one do not show a wave-beating pattern. As documented by microcinematography (Erlandsen and Feely, 1984), and videomicroscopy (Campanati et al., 2002; Ghosh et al., 2001),
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the tail displays a lateral rocking movement and dorso-ventral flexion. Campanati et al. (2002) showed that motility of the caudal region was completely independent from the flagellar beating and it was likely to be produced by a microtubular complex located in the caudal portion of the cell. The funis is a poorly studied microtubular structure. It was described by Holberton (1973) as short arrays of microtubules emanating from the axonemes of the caudal flagella, one in the dorsal and one in the ventral position, and was designated as funis by Kulda and Noh ynkov a (1978). In the internuclei area the funis consists of well-defined bands of interconnected microtubules and from the point of emergence of the ventral flagella, the individual microtubules detach from each band and fan out laterally in a position dorsal to the posterior-lateral axonemes, down to the tail of the organism (Campanati et al., 2003; Kulda and Noh ynkova, 1978). It has been suggested that the funis would have a structural function (Kulda and Noh ynkov a, 1995). The aim of this paper is to describe the fine structure of the funis, its connections, and its possible functional role. High-resolution scanning electron microscopy images of the funis showed that its microtubules do not end in the cytoplasm, but are anchored in the posterior flagella, via the dense rods. In addition, after the emergence of the posterior-lateral flagella, the funis microtubules irradiate towards the plasma membrane and attach to filamentous links underlying the membrane. We suggest that the funis participates on the lateral tail flexion and propose a model for the caudal motility of G. lamblia.
2. Materials and methods 2.1. Organisms and culture Giardia lamblia WB strain (American Type Culture Collection, No. 30957) was cultivated in TYI-S-33 medium enriched with 10% heat-inactivated bovine serum (Diamond et al., 1978) at pH 7.05, without added vitamins, iron, or antibiotics (Gillin et al., 1989), but supplemented with 0.1% bovine bile (Keister, 1983) for 48–72 h, at 37 °C.
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OsO4 in cacodylate buffer containing 5 mM CaCl2 and 0.8% potassium ferricyanide in the dark, for 30 min. Cells were washed in PBS, dehydrated in acetone, and embedded in Epon. Ultra-thin sections were stained with uranyl acetate for 20 min and lead citrate for 5 min and observed in a JEOL 1210 or a Zeiss 900 transmission electron microscope operating at 80 kV. All reagents, unless otherwise stated, were purchased from Sigma, St. Louis, USA. For negative staining PBS washed cells were allowed to adhere to formvar coated grids for 2 min, and the membranes were extracted in a solution containing 1% Nonidet P-40 in PHEM buffer (Schliwa and van Blerkom, 1981) for 40 min. After washes in PHEM buffer, the samples were fixed in 2% glutaraldehyde, washed with water, and then stained with 1% ammonium molibdate for 1 min. Next, the grids were observed in a Zeiss 900 TEM. 2.3. Conventional scanning electron microscopy Living cells were adhered to poly-L -lysine-coated glass coverslips, fixed with 2.5% glutaraldehyde in cacodylate buffer, post-fixed for 5 min in 1% OsO4 , dehydrated in ethanol, critical point dried with CO2 , sputter-coated with gold–palladium, and examined in a Jeol 5800 scanning electron microscope operating at 12 kV. 2.4. High-resolution field emission scanning electron microscopy Cells were adhered to poly-L -lysine coated glass coverslips and then treated with the permeabilization buffer (0.5% Nonidet P-40, 0.1 M Pipes, 1 mM MgSO4 , 2 mM glycerol, 2 mM EGTA, 1 mM PMSF (phenylmethylsulfonyl fluoride), and 0.5% Triton X-100) for different times (10 min–2 h). The cells were washed in PBS and then fixed in 2.5% glutaraldehyde in phosphate buffer, post-fixed for 5 min in 1% OsO4 , dehydrated in ethanol, critical point dried with CO2 , and sputtercoated with carbon. The samples were examined in a Jeol JSM-6340F Field Emission Scanning Electron Microscope operated at an accelerating voltage of 5 kV, using 5 mm as working distance and the standard SEI and BEI detectors.
2.2. Transmission electron microscopy1 2.5. Immunoelectron microscopy Cells were fixed overnight at room temperature in 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2). Then, the cells were washed twice in PBS (phosphatebuffered saline). Post-fixation was performed in 1% 1
Abbreviations used: FESEM, field emission scanning electron microscopy; SEM, scanning electron microscopy; TEM, transmission electron microscopy.
Cells were adhered to poly-L -lysine coated glass coverslips and then treated with the permeabilization buffer as above described. For immunolabeling, the cytoskeletons were first incubated in a solution of PBS containing 3% albumin, quenched in 50 mM NH4 Cl for 30 min, and incubated overnight in the presence of a monoclonal anti-a-tubulin TAT-1 antibody, kindly
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provided by Dr. K. Gull (Woods et al., 1989). After several washes in PBS–1% albumin, the cytoskeletons were incubated in the presence of 15 nm gold-labeled goat anti-mouse IgG (BBInternational, UK). Coverslips were washed several times in PBS and then fixed in 2.5% glutaraldehyde in phosphate buffer, post-fixed for 15 min in 1% OsO4 , dehydrated in ethanol, critical point dried with CO2 , and sputter-coated with carbon. The samples were examined in a Jeol JSM-6340F Field
Emission Scanning Electron Microscope operated at an accelerating voltage of 5 kV, using 5 mm as working distance, and the standard SEI and BEI detectors.
3. Results When observed by routine transmission or scanning electron microscopy Giardia trophozoites showed a
Fig. 1. Routine preparation for transmission electron microscopy of G. lamblia showing the ventral disc (D), the two nuclei (N), peripheral vesicles (V), flagellar axonemes (A), and the funis (arrows). (a) Shows a cross-section where the funis is seen as linear microtubular structures following the caudal flagella (arrows). (b,c) Longitudinal sections showing the funis microtubules (Fn) emanating from the caudal flagella (C). In (c) the brigdes between the funis microtubules are pointed by arrow (inset). MB, median body. Bars ¼ 1 lm.
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half-pear or teardrop shaped body (Figs. 1b, 3a). The ventral disc was seen in the anterior region of the cell and it was laterally and anteriorly surrounded by the marginal groove and the ventrolateral flange (Figs. 1a and 2). Gi-
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ardia presented four pairs of flagella, namely the anterior, posterior-lateral, caudal, and ventral (Figs. 2 and 3a,b). Ultrastructural examination of detergent-treated G. lamblia by FESEM allowed the visualization of its
Fig. 2. Negative staining of G. lamblia using 1% ammonium molibdate. Giardia was extracted by detergents (see Section 2), allowed to adhere on a formvar-coated grid, stained, and observed in the transmission electron microscope. The funis (arrow) is seen, as well the disc (D), and flagella. The nuclei (N) are dorsally situated in relation to the disc. Bar ¼ 1 lm.
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Fig. 3. Routine scanning electron microscopy (a) and field emission scanning electron microscopy (FESEM) after extraction with Triton X-100 (see Section 2) (b–d). In (b) the funis (arrow) is observed as microtubules connecting the axonemes of the caudal flagella (C) to the posterior-lateral flagella (P). The adhesive disc (D) and the four pairs of flagella, the ventral (V), posterior-lateral (P), anterior (A), and caudal (C) are seen. In (c) the caudal region shows the funis microtubules (Fn) emanating from the caudal flagella (C) to the posterior-lateral flagella (P). (c) A higher magnification of the caudal region seen in (b). Note that the funis microtubules originating from the right caudal flagellum cover the left caudal flagellum before reaching the fibrillar material. After the emergence of the posterior-lateral flagella the microtubules are directly connected to the fibrillar material underlying the plasma membrane. In (d), a close view of the caudal flagella (C) from where the funis microtubules (Fn) fan out towards the posteriorlateral flagella (P) and are seen anchored to the dense rods (arrows). Note some of the filamentous bridges interconnecting the funis microtubules. Bars ¼ 1 lm.
microtubular cytoskeleton (Fig. 2). The adhesive disc, located in the ventral anterior region of the trophozoite, was mainly made of concentrically arranged microtubules (Figs. 1a, 2, 3a, 4a, and 5). The disc microtubules and their associated dorsal ribbons were organized in a spiralized organization, surrounding a central area devoid of microtubules, known as the bare area (Figs. 1a, 3a, and 4a). The thickness of the microtubules observed in our preparations varied as a function of the carbon layer thickness used in the sample preparation and also by a coat of fibrous material present in some microtubular structures. This can be compared observing the microtubules of the funis, and the median body (Fig. 6b).
The axonemes of the caudal flagella present emanating microtubules (Figs. 1b,c, 2, 3b–d, and 4–8), which are part of a structure named funis by Kulda and Nohynkova (1978). At the most anterior region of the caudal flagella, near the basal bodies, the microtubules of the funis wrapped the caudal axonemes (Figs. 3d and 8a). Gradually, these microtubules begin to leave the caudal axonemes and fan out towards the axonemes of the posterior-lateral flagella (Figs. 3b–d, 4a,b, and 5–7). These microtubules were interconnected by bridges and attached to the fibrous rod (dense rod) present on the axoneme of the posterior-lateral flagella (Figs. 3d and 6b). Thus, the caudal and posterior-lateral axonemes flagella become linked by the funis microtubules (Figs.
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Fig. 4. High-resolution field emission scanning electron microscopy of Giardia cytoskeleton showing microtubules emanating from the caudal flagella towards the plasma membrane (arrow). Adhesive disc (D); funis (Fn); anterior flagella (A); posterior-lateral flagella (P); ventral flagella (V); and caudal flagella (C). (b) A close view of the ventral region showing the funis microtubules (Fn) which are linked by filamentous bridges (arrow). Posterior-lateral flagella (P); caudal flagella (C); ventral flagella (V); and ventral disc (D). Bars ¼ 1 lm.
3b,c, 5, and 6). When Giardia was adhered through its ventral region (adhesive disc down) on poly-L -lysine coated coverslips, the funis was observed on its dorsal face (Fig. 5). This view allowed the observation that the funis microtubules arised from one of the caudal flagella axonemes, running towards the left, contacting the other caudal flagellum axoneme and adhering to the dense rods of the posterior-lateral flagellum axoneme (Fig. 5). When Giardia was adhered through its dorsal region, (adhesive disc up) it was possible to observe that the other caudal flagellum axoneme also presented emanating microtubules, which are directed to the opposite posterior-lateral flagellar axoneme (Figs. 3, 4, and 6). Thus, there are two emanating groups of microtubules from the axonemes of the caudal flagella that migrate in opposite directions, forming a well-structured array (Figs. 3–6, and 10). In addition, the funis microtubules were covered by an unidentified material, which gave them a greater thickness. When cytoskeletons were isolated by suspending cells in Triton without first attaching them to a surface, several structures, such as microtubules, bridges, and the median bodies were normally lost into the supernatant. Crossley et al. (1986) have made a similar observation. Thus, several trials were made using different times and
detergent concentrations until a good preservation of the bridges was achieved. Only in cells well extracted by detergent the funis were clearly seen. The microtubules of the funis presented bridges (Figs. 1c, 3d, and 6b), which were very sensitive to Triton X-100, and were easily lost during cytoskeleton preparations. By means of the funis microtubules the caudal flagella axonemes seems to be firmly anchored to the axoneme of the posterior-lateral flagella. In addition, after the emergence of the posterior-lateral flagella, few microtubules were seen projecting towards the plasma membrane. They were anchored to a network found underlying the membrane (Fig. 3d). These structures were preserved during sample preparation, using different times of extraction and detergent concentrations, although the bridges interconnecting the funis microtubules were affected when longer times, high detergent concentrations, or centrifugation were used (Fig. 3). The microtubules of the median body were dorsally positioned in relation to the funis and were seen as smooth cylinders, rather distinct from those of the funis, which are covered by a rough coat (Fig. 6). A linearly aligned material, forming images similar to ‘‘corn kernels,’’ covered the posterior-lateral flagella. This coating material was rough and the grains were
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Fig. 5. Scanning electron microscopy showing a dorsal view of the funis microtubules (Fn). They emanate from each of the caudal flagellum (C), contact the other flagellum (arrow) and attach to the dense rod of the opposite posterior-lateral flagellum (P). D, disc; A, anterior flagellum. Note that the arrow and the arrowhead point to funis microtubules found in distinct levels. Bar ¼ 1 lm.
seen in linear rows (Fig. 7). Immunocytochemistry was performed after the detergent extraction and monoclonal anti-tubulin TAT-1, was used (Fig. 8). The funis
microtubules were intensely labeled and were better visualized when back-scattered electrons were used (BEI), as shown in Fig. 8b.
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Fig. 6. (a) SEM of G. lamblia in a ventral view. Note that the microtubules of the funis (Fn) emanate from the caudal flagella (C) and are anchored to the posterior-lateral flagella (P). The median body (MB) is also observed as bundles of microtubules close to the funis. The arrows point to nuclei prints which are located dorsally to the ventral adhesive disc (D). Bar ¼ 1 lm. (b) FESEM of G. lamblia in a close view. The funis microtubules (Fn) are observed emanating from the caudal flagella (C) towards the posterior-lateral flagella (P). Notice that the microtubules present links (arrows), many of which were disrupted by the extraction treatment. The median body (MB), also formed by microtubules, is seen in a dorsal position in relation to the funis. Ventral flagella (V). Bar ¼ 100 nm.
After the emergence of the posterior-lateral flagella, the funis microtubules contacted the plasma membrane (Figs. 3b,c and 4a), via a fibrilar material that underlies the membrane. It is well-known that Giardia presents a tail movement (Fig. 9). When cells are well-fixed the caudal region can be seen in a bent position, either by routine SEM (Fig. 9a) or after detergent extraction (Fig. 9b). We propose that the funis microtubules play some role on this type of movement, as suggested in a schematic drawing (Fig. 10).
4. Discussion Giardia has a complex cytoskeleton formed by different groups of structures that participate in important cell functions, such as adhesion and motility. Although many researchers have investigated motility and attachment processes in Giardia, findings concerning the role of the cytoskeleton in several processes are scant or contradictory. In addition, two components of the in Giardia cytoskeleton received almost no attention: the funis and the median body. Herein we show by the first time images of the cytoskeleton of G. lamblia as visualized by scanning electron
microscopy, allowing to get new insights into several structures. We made use of high-resolution field emission scanning electron microscopy (FESEM) and conventional SEM associated to plasma membrane extraction using different detergent concentrations, and immunogold labeling in order to get #new views of Giardia ultrastructure. Different times, types and detergents concentrations, as well several details on the centrifugation steps were tried before to achieve these preparations. The caudal microtubular structure termed funis (Kulda and Nohynkova, 1978) was carefully analyzed. The funis was described in the literature as a girdle of parallel single tubules that partly enclose the caudal axonemes (Crossley et al., 1986), and as sheets of microtubules (Campanati et al., 2002; Kulda and Nohynkova, 1995). It was considered that the funis microtubules provided a set of lateral ribs reaching the posterior cytoplasm (Campanati et al., 2003; Crossley et al., 1986; Kulda and Nohynkova, 1995). Here we confirmed that the funis is in fact made of microtubules, since they were labeled with anti-tubulin monoclonal antibodies. In addition, we show by the first time the connections between the funis microtubules and the posterior-lateral flagella, suggesting this participation on
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Fig. 7. High-resolution field emission scanning electron microscopy of G. lamblia in high magnification. The funis microtubules (Fn) are clearly seen anchored to the posterior-lateral flagella (P). Adhesive disc (D); ventral flagella (V). Bar ¼ 1 lm.
the Giardia caudal movement. All previous studies proposed that the funis did not present any structural connection when they emerge from the caudal flagella (Crossley et al., 1986; Kulda and Noh ynkov a, 1978, 1995). FESEM observations of the funis showed that this structure is formed by several single interconnected microtubules linking the caudal to the posterior-lateral flagella. Lateral microtubules emanate from each caudal flagellum, contact the other caudal flagellum and attach to the dense rod of the opposite posterior-lateral flagellum. The dorsal group of microtubules arises from the left caudal axoneme, close to the basal body, crosses to the right side of the cell, whereas the ventral group arises on the right, and reaches the left side. Each group of the funis microtubules runs in opposite directions and thus seems to exert more resistance to stress conditions. After the emergence of the posteriorlateral flagella the funis microtubules are seen anchored to the underlying material in the plasma membrane. Many protists posses a unique cortical cytoplasm: the epiplasm, which is a prominent proteinaceous layer
underlying the plasma membrane (Huttenlauch and Stick, 2003). Thus, the funis microtubules seem to be anchored to the epiplasm in the most posterior region of Giardia. It is this region that presents a lateral tail movement. By using our new protocol, new information were obtained on Giardia cytoskeleton. The funis microtubules are interconnected by bridges and coated by an undefined material. On the other hand, one of the microtubules ends is not freely dispersed in the cytoplasm, as proposed in previous studies, but firmly connected to posterior-lateral flagella or to the plasma membrane, after the emergence of the flagella. There are numerous examples in the literature of bridges connecting microtubules (McIntosch, 1974), which are important for maintaining the order of the microtubule array. In several cases such association is involved in the generation of motility. Motor proteins, which power cytoskeletal movements, have been found in other organisms. However, microtubule-based motor proteins, such as kinesin and dynein, and the microfilament-based myosin, are virtually uncharacterized in
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Fig. 8. High-resolution field emission scanning electron microscopy of G. lamblia after immunogold labeling using the monoclonal antibody TAT-1. In (a), the image is seen by SEI (a) whereas in (b) the same cell is shown by BEI. It is possible to see the gold particles labeling the microtubules of the adhesive disc (D), funis (Fn), caudal (C), and posterior-lateral flagella (P). Bar ¼ 1 lm.
Giardia (Elmendorf et al., 2003). Campanati et al. (2002) using high-pressure freezing and freeze-substitution observed arms emanating from the funis micro-
tubules. They assumed that the tiny filaments attached to the microtubules of these sheets might be dynein molecules. However, immunocytochemistry location of
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Fig. 9. Conventional scanning electron microscopy of G. lamblia seen on dorsal view, without membrane extraction (a) and in ventral view, after membrane extraction by detergents (b). Notice the caudal bent. D, disc; P, posterior-lateral flagella; C, caudal flagella; A, anterior flagella; V, ventral flagella; and Fn, funis. Bars ¼ 1 lm.
this protein using commercial antibodies was unsuccessful. In the present report, we observed these arms more clearly, and they were also contacting the neighbor microtubules.
The parasite motion is complex. Giardia displays different types of movements either when swimming or attached. The four pairs of flagella control different aspects of motility (Elmendorf et al., 2003). While the
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Fig. 10. Schematic drawing of a static G. lamblia (a,b) and presenting tail movement (c) showing the relationship of the funis and the posterior-lateral and caudal flagella. The caudal flagella pair (C1,C2) present an array of microtubules (in lilac and green) that emanate towards the posterior-lateral flagella (P1,P2). Note that in the caudal flagellum C2 the microtubules emanate towards the dense rods of the posterior-lateral flagellum 1 (P1), whereas the caudal flagellum 1 (C1) the microtubules are directed towards the posterior flagellum 2 (P2). Notice that the funis microtubules are anchored to the dense rods (in red) of the posterior-lateral flagella, and are interconnected by filamentous bridges (orange). After the emergence of the posterior-lateral flagella the funis microtubules are anchored to the epiplasm, a layer of material underlying the plasma membrane. This region presents a network of filamentous structures (F). The pair of caudal flagella is interconnected by thin bridges. During tail flexion (c) the microtubules of the funis found after the posterior-lateral flagella emergence are seen anchored to the material underlying the plasma membrane and presenting different length. They seem to participate in the caudal movement. D, disc; PV, peripheral vesicles. Bars ¼ 1 lm.
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parasite is attached, the ventral flagella beat continuously and when the cells are freely moving they present either a lateral rocking or a complete rotational (tumbling) movement along its longitudinal axis (Campanati et al., 2002; Elmendorf et al., 2003). An interesting phenomenon observed in G. lamblia is the movement performed by its tail region. This portion of the cell is able to bend both laterally and/or dorsoventrally. This movement is completely independent from the flagellar beating (Campanati et al., 2002; Glesbki, 1967). It was proposed that the microtubular complex located in the caudal portion of the cell likely produced the dislocation of the caudal region (Campanati et al., 2002). However, even performing highpressure freezing of living cells and freeze-substitution, structural details of this caudal region were not achieved in previous studies (Campanati et al., 2002). In the present study, FESEM associated with a detergent extraction protocol, allowed a better understanding of this cell region. The mechanism of dorsal and lateral flexion is not completely understood. It was suggested that this flexion is structurally associated in some way with the caudal flagella (Campanati et al., 2002), but it remains to be determined whether this force is derived only from flagellar movement or also from contractile proteins, associated in some fashion with the flagellar axonemes or the funis (Erlandsen and Feely, 1984). The structural organization of funis microtubules observed in this study perfectly fits with the observations made by Campanati et al. (2002) using video-microscopy in which the caudal flagella axonemes were seen rolling over each other, during the lateral dislocation of the caudal region of the cell. Dense fibrillar material has been reported by Feely et al. (1982) to parallel the course of the caudal flagella and their accompanying microtubules (funis), but the nature of this fibrillar material is unknown. Feely et al. (1982) found actin and a-actinin staining by immunofluorescence microscopy in the region corresponding to the dense rod-like masses near the axonemes of the posterior-lateral flagella and Narcisi et al. (1994) found actin along the intracellular axonemes. Feely et al. (1982) raised the possibility that the association of contractile proteins and microtubules could have functional significance. Careful observation of their images suggested us an intense labeling in the area of the funis. This finding, if confirmed, could explain the caudal contractility. Contractile proteins could participate, together with the funis microtubules, in the tail flexion. We observed that both the caudal and posterior-lateral flagella present a coat made of an uncharacterized material. The posterior-lateral flagella presented this dense coat covering their axonemes, whereas in the caudal flagella the coat covers only some areas, probably the regions of higher stress. In addition, a dense material
also covers the caudal flagella. It is necessary to determine the nature of the proteins composing these dense rods. Campanati et al. (2002) observed by video-microscopy that during the lateral movement of the caudal region one axoneme seemed to roll over the other one. They proposed that the movement generated by the inner axonemal apparatus could be constrained by the accessory microtubular structures and by the other axoneme, producing the movement of the caudal region. Our observations showed that indeed one group of microtubules originated from one of the caudal flagellum running over the other flagellum and in this course established contact with the other caudal flagellum (see scheme in Fig. 10). Like the interaction of microtubules in the axoneme, a sliding of one microtubule over the other could lead to shortening of the funis and, as consequence, the tail movement. We propose that the funis works as a stress and flexible cord. Thus, the name funis proposed by Kulda and Nohynkova (1978) is adequate (it means rope, cord). We speculate that the funis could be responsible for the caudal lateral movement in G. lamblia. This could be accomplished by the sliding of a microtubule over the other, as occurs in movements of flagellar axonemes. In addition, contractile proteins associated with funis microtubules could also participate in this movement. Another hypothesis includes a rapid polymerization/ depolymerization of microtubules in the region of insertion to the dense rods in the posterior-lateral flagellum, leading to caudal lateral flexion in Giardia.
Acknowledgments This work was supported by the Conselho Nacional de Desenvolvimento Cientıfico e Tecnol ogico (CNPq), Fundacß~ao Carlos Chagas Filho de Amparo a Pesquisa do Estado do Rio de Janeiro (FAPERJ), Programa de N ucleos de Excel^encia (PRONEX), Coordenacß~ ao de Aperfeicßoamento de Pessoal de Nıvel Superior (CAPES), and Associacß~ao Universitaria Santa Ursula (AUSU). The authors thank the technical support of William Christian Mol^edo Lopes.
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