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Identification and organization of the cytoskeleton in the alga Vaucheria longicaulis var. macounii Peat, Lucinda Jane 1992

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IDENTIFICATION AND ORGANIZATION OF THE CYTOSKELETON IN THE ALGA Vaucheria lonqicaulis var. macounii by LUCINDA JANE PEAT B.Sc, The University of Leeds, U.K., 1986 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF THE FACULTY OF GRADUATE STUDIES (Department of Botany) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA September 1992 (c)Lucinda Jane Peat, 1992 MASTER OF SCIENCE in In presenting this thesis in partial fulfiinnent of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. 1 further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives, it is understood that copying or publication of this thesis for financial gain shall not be allowed without my wri t ten permission. Department of The University of British Columbia Vancouver, Canada Date DE-6 (2/88) Abstract The presence of the cytoskeletal proteins actin and tubulin in the alga Vaucheria lonqicaulis Hoppaugh var. macounii Blum i s studied by SDS-PAGE and immunoblotting techniques. These techniques also indicate the presence of the mechanomotor protein myosin. The overall organization of the cytoskeleton in the cytoplasm of intact filaments, their identification and distribution, are investigated by immunofluorescence and epifluorescence microscopy using monoclonal anti-B-tubulin and anti-actin antibodies and FITC-labelled phalloidin. ' Anti-myosin antibodies were also utilized, but proved to be inadequate for my work. Phalloidin labelling of F-actin proved to be more suitable for visualizing microfilaments than anti-actin antibodies. Phalloidin labelling of F-actin reveals a dense array of microfilament cables in the cortical cytoplasm of vegetative filaments, which appear to be sub-divided into two morphologically distinct sets. One set consists of straighter elements, preferentially occupying the cytoplasm adjacent to the plasma membrane and possibly providing tracks for organelle motility. A second set i s made up of wavy elements and extends deeper into the cytoplasm where i t may be part of the force generating system responsible for organelle motility. Immunofluorescence for tubulin reveals that the microtubule array i s much less dense than that made up of microfilaments. Microtubule bundles appear shorter and straighter and are located throughout the width of the cytoplasm. They show no particular relationship to organelles, except nuclei. With respect to nuclei, they seem to be involved in their arrangement within the filament, particularly in the apical region, and this may have implications for the organization of the tip-growth processes. High resolution scanning electron microscopy i s also u t i l i z e d in the study of the organization and distribution of the cytoskeleton. Differential interference contrast microscopy reveals cytoplasmic tracks originating from focal regions in l i v i n g vegetative filaments of the coenocytic alga Vaucheria longicaulis var. macounii. Fluorescein-labelled phalloidin staining also reveals regions (foci) of similar structure and dimensions among the F-actin array. Immunofluorescence using monoclonal antibodies to B-tubulin shows punctate fluorescence in association with the microtubule array. Cytochalasins are used to breakdown the F-actin array, an effect that i s concentration dependent. Cytochalasin D causes a gradual breakdown of the F-actin array, revealing the close association between foc i and F-actin fluorescence. Recovery from treatment with this inhibitor confirms this association and suggests that foci act as organizing centers for the F-actin array. Cold temperature, Oryzalin and taxol are used to disrupt the microtubule bundle array. Depolymerization is evident in the appearance of many fluorescent spots (foci), which may be co-localized with nuclei or associated with the ends of microtubule bundles. Recovery from these treatments suggests that spot-like foci act as nucleation centers for microtubule bundles. The existence of microtubule-associated foci i s supported by the results of taxol treatment. The different roles played by microfilaments and microtubules in the organization of the polarized structure of the c e l l are discussed with respect to the function of their respective organizing centers. TABLE OF CONTENTS ABSTRACT i i TABLE OF CONTENTS V LIST OF FIGURES v i ACKNOWLEDGEMENTS x i INTRODUCTION 1 MATERIALS AND METHODS 6 RESULTS - PART 1 12 PROTEIN EXTRACTION AND ANALYSIS 12 DIFFERENTIAL INTERFERENCE MICROSCOPY OBSERVATIONS 13 MICROFILAMENT ARRAY AS VISUALIZED BY FLUORESCENCE 13 MICROTUBULE ARRAY AS VISUALIZED BY IMMUNOFLUORESCENCE..16 HIGH RESOLUTION ELECTRON MICROSCOPY OBSERVATIONS 18 DISCUSSION - PART 1 19 RESULTS - PART 2 28 FOCAL REGIONS IN UNTREATED CELLS 28 F-ACTIN FOCI 29 MICROTUBULE ASSOCIATED FOCI 32 DISCUSSION - PART 2 35 CONCLUSIONS 41 FIGURES 43 LITERATURE CITED 59 LIST OF FIGURES FIGURE 1 SDS-PAGE gels and immunoblots of Vaucheria homogenate proteins 44 2 DIG image of axially aligned array of fibrous structures distributed throughout the cytoplasm 45 3 DIG image showing relationship between fibrous structures,chloroplasts, and nuclei 45 4 F-actin microfilament array visualized using fluorescein-labelled phalloidin. inset: distribution of F-actin in cortical region of the filament 45 5 Microfilament array showing branches between longitudinal cables and around organelles 45 6 Branching of microfilament cables in close proximity to organelles 46 7 Grystalline-like microfilament arrays in Vaucheria filament. 46 8 Association of microfilament cables and chloroplasts...47 9 Microfilament cables appear to radiate from a focal area in the apical region of a vegetative filament 47 10 Microfilament cable organization into two sub-sets: a straighter sub-set, and a more reticulate one 47 11 Focal point of microfilament cables ......47 12 Confocal scans showing the partitioning of the microfilament array 48 13 Confocal scans showing a longitudinally aligned microfilament set and a reticulate set beneath i t 48 14 Confocal cross-sectional scan of microfilament distribution within the cytoplasm 48 15 Confocal graphical representation of microfilament distribution along a labelled longitudinal axis of the c e l l 48 16 Microtubule bundle array of a Vaucheria vegetative filament 49 17 Vaucheria c e l l showing microtubule bundles curving and crossing over into a branch 49 18 Confocal composite image of microtubule bundles 49 19 Microfilament bundle array in the t i p region of the filament 49 20 Cross-sectional confocal scan and graphical representation of microtubule bundle distribution throughout the cytoplasm 50 21 Serial confocal scans at different optical depths showing microtubule bundles throughout the cytoplasm...50 22 Cross-section of a Vaucheria filament showing punctate microtubule fluorescence throughout the cytoplasm 50 2 3 Extruded chloroplast with closely associated microtubule bundles ......50 24 HRSEM of a fractured end of a Vaucheria c e l l . . . 51 25 Higher magnification HRSEM of fibrous material 51 26 HRSEM of a fractured filament showing straighter cables within the cytoplasm 51 27 Higher magnification of Fig. 26 51 2.1 DIG image showing cytoplasmic track foci 53 2.2 Microfilament array showing foci adjacent to a cross -wall 53 2.3 Microtubule array of an untreated (control) c e l l 53 2.4 Oblique end view of a vegetative filament revealing filamentous and punctate B-tubulin fluorescence 53 2.5 F-actin fluorescence after treatment with Gytochalasin D 54 2.6 F-actin fo c i after treatment with Cytochalasin B 54 2.7 Early stage of F-actin breakdown by Cytochalasin D 54 2.8 F-actin array of an untreated vegetative filament 54 2.9 F-actin foci in the vicinity of a cross-wall after Gytochalasin D treatment 54 2.10 F-actin fo c i amidst F-actin cables after treatment with Gytochalasin D 54 2.11 Web-like array of F-actin with foci after treatment with Cytochalasin D 55 2.12 Web-like array of F-actin with foci after treatment with Gytochalasin D - tip region of vegetative filament 55 2.13 Advanced stage of F-actin depolymerization after treatment with Cytochalasin D 55 2.14 Extensive depolymerization of F-actin after treatment with Cytochalasin D 55 2.15 Almost complete F-actin depolymerization after treatment with Cytochalasin D 55 2.16 F-actin array after cold treatment - appearance of foci 55 2.17 Foci during recovery from Cytochalasin D treatment 56 2.18 Microtubule array after cold treatment 56 2.19 Depolymerization of the microtubule array after cold treatment 56 2.20 Depolymerization of the microtubule array after cold treatment - t i p region 56 2.21 Distribution of B-tubulin fluorescence after cold treatment 56 2.22 DAPI localization of nuclei as a reference to Figure 2.21 56 2.23 Fragmentation of microtubule bundles during depolymerization 57 2.24 Bright spots at ends of depolymerizing microtubule bundles 57 2.25 Bright spot at end of depolymerizing microtubule bundle 57 2.26 Microtubule bundle array after treatment with taxol....57 2.27 Fluorescent spots visi b l e during repolymerization of the microtubule array ......58 2.28 Microtubule bundles during the process of repolymerization 58 2.29 Fluorescent spot terminated microtubule bundles during repolymerization 58 2.30 Repolymerization of the microtubule array 58 ACKNOWLEDGEMENTS It has been an enriching experience to study and live abroad. I would like to express thanks to those who made this possible; my supervisor Dr. Luis Oliveira, UBC, Canada Immigration, and my family and friends on both sides of the world for their unceasing support of my endeavours. Special thanks are in order for technical advice and assistance to Dr. Brian Oates and Dr. Edith Camm and her laboratory staff at the Department of Botany, UBC, and Dr. John W. La Claire II at the University of Texas, Austin, for performing the anti-Dictyostelium myosin immunoblot. Introduction The cytoskeleton of higher plant and algal c e l l s i s , for the most part, considered to be made up of microfilaments and microtubules (Seagull 1989, Derksen et a l . 1990), although recently intermediate filament-like structures have been described (Goodbody et a l . 1989). Multiple functions have been assigned to these cytoskeletal elements. They include intracellular motility (Williamson 1986), roles in mitosis and cytokinesis (Forer 1982, Gunning 1982), spatial organization (Van Lammeren 1988), maintenance of c e l l shape (Marchant 1982) and alignment of c e l l wall microfibrils (Robinson and Quader 1982). The involvement of the cytoskeleton in intracellular motility and the creation of motive force have been studied in depth using giant algal cells as models, e.g. Ghara (Williamson 1975, Grolig et a l . 1988). In such c e l l s i t has been determined that the major mechanism underlying these processes u t i l i z e s an actomyosin system. In many higher plant and algal c e l l s exhibiting cytoplasmic streaming, extensive axially aligned arrays of actin microfilaments can be seen using immunofluorescence and fluorochrome labelled phallotoxins (Menzel 1987, Heslop-Harrison and Heslop-Harrison 1989). They are often presumed to function in a similar fashion to the algal c e l l models. In contrast, microtubule based motive systems which have been found to be largely responsible for intracellular motility within animal c e l l s (Adams and Pollard 1986), where they seem to participate in this function by acting in conjunction with mechanoenzymes such as dynein, kinesin and other microtubule associated proteins (Hollenbeck 1988). In streaming higher plant and algal c e l l s immunofluorescence staining of tubulin has revealed microtubule arrays which are sometimes axially co-aligned with F-actin arrays. As a result of inhibitor studies and molecular analysis, they have also been implicated to function in the movement of organelles within c e l l s (Menzel and Schliwa 1986b, Schnepf 1986, Tiezzi et a l . 1992, Tornbom and Oliveira, unpublished). Tip growing ce l l s , examples of which include pollen tubes (Steer and Steer 1989), moss protonemata (Sievers and Schnepf 1981), algal rhizoids (Tewinkel et a l . 1989) and fungal hyphae (Temperli et a l . 1990), demonstrate a unique property of localized growth and zonation of organelles as a manifestation of polarization (Sievers and Schnepf 1981, Schnepf 1986). Cytoskeleton has also been found to be integral in the establishment and maintenance of t i p growth. In such c e l l s microtubules and microfilaments are also usually axially aligned (Lloyd 1987) and extend into the apical dome of the growing tip where they are implicated to function in the maintenance of structural integrity as well as in the processes of exocytosis and streaming (Doonan et a l . 1988, Steer and Steer 1989). Vaucheria lonqicaulis Hoppaugh var. macounii Blum is a coenocytic filamentous alga of the Division Chrysophyta, which exhibits t i p growth in the apical dome of each filament (Kataoka, 1975). Rapid cytoplasmic streaming of organelles, including chloroplasts, nuclei, mitochondria and small vesicles (Ott 1992, Tornbom and Oliveira, unpublished), i s also observed throughout the body of the filament into the t i p region. Previous studies have revealed the presence of cytoskeletal elements in Vaucheria (Ott and Brown 1972, Blatt et a l . 1980, Oliveira and Fitch 1988, Ott 1992, Tornbom and Oliveira, unpublished) but these did not include the use of fluorescence microscopy and protein analysis. These techniques are used in this study in order to elucidate the organization of, and role, the cytoskeleton plays in this unusual c e l l type with complex intra-cellular motility and polarized growth (Tornbom and Oliveira, unpublished). Since the recognition of the cytoskeleton as an ubiquitous component of eukaryotic cells, the subject of how the individual proteinaceous elements are organized and assembled has been of great interest. Observations of how inhibitors disrupt (depolymerize) cytoskeletal elements can reveal information such as the existence of microtubule organizing centers (MTOCs) (Brown et a l . 1982) or centers of actin assembly (polymerization) (Stossel 1989). The organization of filamentous actin (F-actin) in plant cel l s has been the subject of numerous studies (Staiger and Schliwa 1987, Seagull et a l . 1987). Despite this fact, our knowledge of the subject is meager compared to that of F-actin in animal c e l l s . In animal cells i t has been shown that actin, actin binding proteins and the plasma membrane are closely associated with one another (Luna 1991, Isenberg 1991). Depolymerization experiments with the fungal metabolites Cytochalasins, which specifically bind to F-actin (Brenner and Korn 1980, Goddette and Frieden 1986), show varied effects often including fragmentation and the formation of foci amidst actin microfilaments. As a result of their structure and behaviour, foci have been described as centers of F-actin organization (Schliwa 1982, Yahara et a l . 1982). Repolymerizing arrays of F-actin arising from these focal regions give supporting evidence for their participation in F-actin assembly and provide information on how the organization of the microfilament network and i t s association with organelles i s established (Parthasarathy 1985). Polymerization of tubulin in plant c e l l s i s observed in conjunction with nucleation sites (Hogetsu 1986). However, the occurrence of highly structured microtubule organizing centers (MTOCs) i s less common in higher plant than animal c e l l s (Lloyd 1987). Possible sites for nucleation of microtubules include the nuclear envelope (Wick and Duniec 1983, Clayton et a l . 1985, Lloyd 1987, Falconer et a l . 1988, Marc and Palevitz 1990, Astrom et a l . 1991), regions of the cortical cytoplasm (Hogetsu 1986, Cleary and Hardham 1990, Marc and Palevitz 1990) and the plasma membrane (Falconer et a l . 1988, Marc and Palevitz 1990, Astrom et a l . 1991). Microtubule inhibitors and low temperatures can be used to disrupt the organization of the microtubular system and often to depolymerize microtubules completely. Repolymerization experiments can then give insight into the establishment of microtubule growth, the pattern of orientation and interaction with different cellular compartments, particularly nucleation sites (Mitchison and Kirschner 1984, Murray 1984, Bre et a l . 1987) . In this study Cytochalasins are used to investigate the involvement of microfilament foci in the assembly and organization of the F-actin array. The microtubule depolymerizing herbicide Oryzalin (Strachan and Hess 1983, Cleary and Hardham 1988 & 1990, Wasteneys and Williamson 1989), the stabilizing drug taxol (Horwitz 1992), and cold temperature which depolymerizes some microtubule systems (Troutt et a l . 1990, Akashi and Shibaoka 1991, Astrom et a l . 1991), are also used to investigate the assembly and organization of the microtubule bundle array and to determine whether or not the B-tubulin spots act as nucleation centers. Materials and Methods Culture procedures Vaucheria lonqicaulis var. macounii was collected from an intertidal region of North Vancouver, B.C., and grown on i t s natural substrate, supplemented with half strength Instant Ocean (Aquarium Systems, Inc., Eastlake, Ohio) medium with added minor elements (Lewin 1966) and s o i l extract. Cultures were kept at 10 °C, with v i t a - l i t e fluorescent lighting providing 250 /xE m~^  s~-^ under a 16-8h light-dark photoperiod. Differential interference contrast and fluorescence microscopy Filaments of V^ lonqicaulis var. macounii selected for microscopical studies were removed from actively growing cultures. Detailed observations and fluorescence micrographs were obtained using a Leitz Dialux 20EB compound light microscope equipped with epifluorescence optics, and a Zeiss Axiophot microscope equipped with differential contrast optics. Confocal laser scanning microscopy images were obtained from a BioRad apparatus with attached Zeiss Axiophot microscope. Fluorescein labelled phalloidin (Sigma Chemical Co, USA) stock solution was made up in 100% methanol at 2 x 10~^ M and stored at -20 °C. The stain was reconstituted in 0.2 M phosphate buffered saline (PBS) (osmolarity equal to that of the growth medium) containing 10 mM EGTA (ethylene glycol-bis (B-aminoethyl ether) N,N,N,N'-tetraacetic acid) and 1 mM MgS04 (and 2%'ethanol or 1% dimethylsulphoxide (DMSO) for permeabilization purposes (Pierson 1988, Heslop-Harrison and Heslop-Harrison 1991). Incubations were carried out for 15 minutes at room temperature. For controls, competition staining was carried out by pre-incubation with unlabelled phalloidin at 1.2 x 10"-^  M before incubation with fluorescein-labelled phalloidin, as previously described. Cells were rinsed in PBS before examination., For indirect immunofluorescence, ce l l s were fixed using freshly made 3% formaldehyde, 0.1% glutaraldehyde in PBS or cytoskeleton stabilizing buffer (5 mM MgS04, 100 mM PIPES (piperazine-N,N'-bis [2-ethanesulphonic acid]), 10 mM EGTA pH 7.3) for 1 hour at room temperature or 4 °C. Use of sodium borohydride after fixation did not vis i b l y improve the fluorescence image. Fixed cells were placed on a poly-L-lysine (Sigma Co., USA, > 300,000 MW) coated slide, in a drop of PBS and cut into small fragments (1 mm or shorter). The PBS was then removed and replaced with 10% skim milk as a blocking agent against non-specific antibody binding. Primary antibodies were monoclonal mouse anti-B-tubulin at dilution 1:300 - 1:500, monoclonal mouse anti-actin at dilution 1:1000. Secondary antibody was goat anti-mouse at dilution 1:30. A l l antibodies were from Amersham Corp., USA. A polyclonal rabbit anti-tubulin (Sigma Co, USA) at dilution 1:10 was used as control. Dilutions were made using PBS containing 0.1% NaN3 and 1 mg/ml BSA (bovine serum albumin) pH 7.3. Staining was improved with the use of 1% DMSO in PBS as a rinsing solution between antibody incubations (Schroeder et a l . 1985, Traas et a l . 1987) . Final mounting medium was a 1:1 solution of glycerol and PBS, containing 0.1% phenylenediamine as an anti-quenching agent. When required, sections of fixed c e l l s were obtained using a freeze stage microtome. High resolution scanning electron microscopy Specimen preparation followed the protocol of Blackmore et a l . (1984), modified for the preservation of cytoskeletal elements. This involved rapid freezing in liquid propane before fracturing the specimens in liquid nitrogen. Specimens were examined using a Hitachi S-4100 high resolution f i e l d emission scanning electron microscope, at an accelerating voltage of 5.0 kV. Electrophoresis and immunoblotting Crude protein extracts of Vaucheria were made by freezing approximately 0.4 g of tissue in liquid nitrogen, and grinding i t in extraètion buffer on ice (La Claire 1991 and references therein). The remaining suspension was centrifuged, the supernatant diluted 1:4 with electrophoresis sample buffer and incubated in boiling water for 5 minutes. Approximately 5 fig of Vaucheria protein per well was loaded (calculated with the BioRad Protein Assay using the Bradford method) before electrophoresis was performed. The bovine brain tubulin was a generous g i f t from Dr. W. Vogl, Dept. Anatomy, UBC, and the chicken gizzard myosin was from Sigma Co, USA. SDS-PAGE (sodium dodecyl sulphate-polyacrylamide gel electrophoresis) was carried out acearding to Laemmli (1970) using a BioRad Protean II apparatus and 16 cm x 0.75 mm thick 12% polyacrylamide gels, except for the myosin immunoblot when a 7.5% gel was used. Semi-dry transfers were performed onto nitrocellulose (Towbin et a l . 1979) which was then blocked using 3% f i s h skin gelatin or 0.5% skim milk (Johnson et al . 1984). Anti-actin and anti-B tubulin monoclonal primary antibodies were as used for immunofluorescence but at a higher dilution (1:1000). Monoclonal anti-pan myosin (Amersham Corp., USA) was used at a dilution of 1:10. The secondary antibody was goat anti-mouse alkaline phosphatase (Gibco BRL, Canada) used at a dilution of 1:1500, and the colour reaction solution contained Fast Red salt and Naphthol AS-MX phosphoric acid (3-hydroxy-2-naphthoic acid-2,4-dimethylanilide phosphate) (Sigma Co, USA) in 50 mM Tris HCl pH 8.0. Affin i t y purified rabbit polyclonal anti-Dictyostelium myosin (Dr. J. W. La Claire, University of Austin, Texas) was diluted 1:250 in Tris buffered saline with 5% milk and 0.5 mg/ml NaN3, and the secondary antibody was goat anti-rabbit alkaline phosphatase (Sigma Co, USA) diluted at 1:1000. Depolymerization experiments Stock solutions of Cytochalasin B and D were made at 20 mg/ml and 1 mg/ml in 100% DMSO, and diluted with growth medium to working concentrations of 200 ^ g/ml and 10 /ig/ml respectively. Stock solution of Oryzalin was made at 7.5 mM in 100% acetone and diluted with growth medium to a working concentration of 10 M M . Stock solution of taxol was made at 2.3 mM in 100% DMSO and diluted with growth medium to 30 - 50 /xM. A l l stock solutions were stored at -20° C. DAPI (4,6-Diamidino-2-phenylindole) was made up directly in growth medium at a concentration of 5 /xg/ml. A l l chemicals were purchased from Sigma Chemical Co., except for Oryzalin [4(Dipropylamino)-3,5-dinitrobenzenesulfonamide, synonym SurfIan] which was a generous g i f t from Dow Elanco Co., Greenfield, Indiana, and taxol, which was a generous g i f t from the National Cancer Institute, Bethesda, Maryland. When appropriate controls using 1% DMSO or 0.1% acetone in growth medium were also carried out, and these solvents revealed no changes in the respective cytoskeletal arrays. Cells were incubated at room temperature for approximately 10 minutes with Cytochalasins and 15 mins with Oryzalin. Cold temperature treatments were performed by cooling vials of growth medium to freezing temperature with a thermoelectric cold plate (Thermoelectrics Unlimited, Inc), the average temperature was -2° C and the cooling period approximately 30 minutes. Temperatures were measured using a calibrated d i g i t a l thermometer probe. Repolymerization experiments After the depolymerization treatments, c e l l s were transferred to inhibitor-free medium, or medium at 10 °C, as appropriate. They were then periodically examined under a microscope for the restoration of cytoplasmic streaming, and stained to assess the degree and sequence of reconstitution of both the microfilament and microtubule arrays. s t a t i s t i c a l analysis Thirty microfilament cables and microtubule bundles were measured from micrographs in order to calculate the mean width, and test whether microtubule bundles are significantly wider than microfilament cables. Means in the text are provided with standard error figures. The difference between microfilament cable and microtubule bundle mean width was found to be significant (p = .002), using the Student t test (Parker 1980) where population variances are not assumed to be equal. However, these measurements are considered to be approximate due to the exaggerated effect of the emitted light from the fluorochrome. Results - Part 1 Protein extraction and analysis Bands of the appropriate molecular weight for actin (-42 kDa) and tubulin (-55 kDa) are consistently present in the t o t a l protein extracts of Vaucheria lonqicaulis var. macounii. In contrast a putative myosin band (-180 Mr) which co-migrates with the myosin molecular weight marker, i s only present upon careful extraction using protease inhibitors and conditions designed to reduce protease activity (Fig. 1). Immunoblots using commercial monoclonal primary antibodies raised to bovine brain tubulin (B sub-unit) and chicken gizzard actin revealed positive cross-reactions with Vaucheria proteins. The anti-actin cross-reacted with only a single band of approximately 47 Mr, whereas the anti-B tubulin cross-reacted with two bands of approximately 55 Mr and 46.5 Mr (Figs. lA & B). Immunoblots using Amersham's monoclonal anti-pan myosin gave a positive cross-reaction with bands of approximately 47 Mr, indicating the presence of probable myosin breakdown products (Fig. IC). Immunoblots were also performed using a polyclonal anti-Dictyostelium myosin antibody, which showed only a very faint indication of cross-reaction at approximately 116 Mr (Fig. ID). Controls for the immunoblotting procedure included omission of the primary antibody (negative control) and use of purified protein standards (positive control). The negative controls showed no cross-reaction, indicating absence of endogenous phosphatase activity, while the positive controls showed cross-reactions with the tubulin and myosin bands, confirming the effectiveness of these antibodies for their identification. Differential interference contrast microscopy observations Differential interference contrast microscopy shows the cytoplasm of vegetative filaments of Vaucheria longicaulis var. macounii to contain an array of cable-like filamentous structures which resemble tracks (Figs. 2 & 3). These are observed throughout the length of the vegetative filaments. The elements of this array appear as long, mostly unbranched cables lying parallel to the longitudinal axis of the c e l l . Their density varies along the length of the c e l l . The organization of the elements of this array i s constantly changing with time. This occurs concurrently with the transport of the organelles they are closely associated with, such as nuclei, small particles and chloroplasts. Microfilament organization; fluorescein conjugated phalloidin studies Figure 4 shows labelling of F-actin using fluorescein-conjugated phalloidin in the mid-filament region of a Vaucheria longicaulis var. macounii c e l l . Controls for this procedure involved examination of the cells after incubation in phalloidin-free solution, or after incubation with high concentration unconjugated phalloidin (in order to saturate the actin binding sites) followed by incubation with fluorescein-conjugated phalloidin. The controls showed l i t t l e or no positive staining (not shown). Actin microfilaments are not easily visualized by indirect immunofluorescence using the Amersham anti-actin antibody, or by phalloidin-FITC staining after conventional aldehyde fixation. However, good preservation of these elements can be achieved by incubating the ce l l s with phalloidin (which binds to actin and stablizes the microfilaments) before aldehyde fixation (Fig. 5). F-actin i s seen to be organized into cable-like strands, which extend throughout the length of the vegetative filaments (Figs. 4 & 5) . Each cable-like strand averages 0.37 ± 0.03 /xm in diameter, and approximately 16 strands seem to occur across the c e l l per f i e l d of view. Taking into consideration that the average thickness of a microfilament i s 5-7 nm (Staiger and Schliwa 1987, Derksen et a l . 1990), each cable-like strand consists of about 60 individual microfilaments. The preponderant orientation of these strands i s parallel to the longitudinal axis of the vegetative filaments. Local branching of microfilament cables occurs in close proximity to organelles (Fig. 6). Occasionally microfilaments were visualised as shorter and more angular crystalline arrays (Fig. 7), consistent with alterations due to osmotic effects on the molecular structure of actin (Janmey 1991). Microfilament strands are often seen to be closely associated with chloroplasts (Fig. 8), forming channel-like regions through which these organelles may travel during cytoplasmic streaming. Towards the t i p region of each vegetative filament, the cable-like strands extend into the apical dome, become more reticulate, and form areas of very intense fluorescence (Fig. 9). When observed from a different perspective a fine network of cables can be seen to radiate from these bright fluorescent areas (Fig. 9, inset). Through multi-level focussing (Fig. 4, inset) and confocal laser scanning microscopy (Figs. 12 & 13), i t i s possible to demonstrate that the whole microfilament array i s almost exclusively located in the f i r s t few microns of the (cortical) cytoplasm (actual depth depending on the individual size of the c e l l ) . Figures 14 and 15 quantify F-actin distribution along the transverse and longitudinal axes of the vegetative filament respectively. The microfilament population i s distributed uniformly both circumferentially (Fig. 14) and longitudinally (Fig. 15) in the c e l l . Microfilament cables closer to the plasma membrane seem straighter and axially aligned, whereas those placed at a greater depth are more reticulate and in closer contact with chloroplasts (Fig. 10). This i s verified by the confocal s e r i a l scans (Fig. 13) which demonstrate reticulate actin cables (3rd and 4th scan) at a greater depth than the axially aligned ones (1st and 2nd scan). A variable number (up to 20) of focal points from which 4 to 10 microfilament cables are seen to radiate occurs in each c e l l . These are found both in the apical region (Fig. 9) as well as throughout the vegetative filament (Fig. 11 & inset). Figure 13 also indicates the existence of foci amidst the reticulate population of microfilaments seen in the 3rd and 4th scans. Microtubule organization: indirect immunofluorescence studies Optimum positive staining of the microtubular compartment was obtained by subjecting fixed material, cut into small fragments, to permeabilization with low concentration DMSO (1%) and subsequent staining with the appropriate antibody solution. Incubation periods of up to 72 hours were necessary to allow diffusion of the antibodies into the cut ends and throughout the body of the c e l l fragment. Controls for this procedure consisted of incubation with only the secondary antibody, or incubation with a primary antibody raised in a different animal (rabbit) to that of the experimental primary antibody (mouse). Both types of control methods resulted in cells with no specific labelling and only dim chloroplast autofluorescence was v i s i b l e (not shown). Figure 16 shows labelling of the B-tubulin in the mid-filament region of a vegetative c e l l . B-tubulin can be seen to be organized into cable-like structures measuring approximately 0.57 + 0.05 jum in diameter, significantly wider than the actin microfilament cables. Assuming that the average microtubule measures 25 nm in diameter (Bershadsky and Vasiliev 1988, Derksen et a l . 1990), these cables consist of approximately 23 individual microtubules. The mean number of microtubule bundles per f i e l d of view i s 7, significantly less than the density of microfilament cables. Microtubule bundles are always straight and shorter (tens of microns in length) than microfilament cables (hundreds of microns). This i s determined through confocal scanning laser microscopy observations where successive optical scans show that microtubule bundles do not span the entire length of the vegetative filament (Fig. 18). The preponderant orientation i s parallel to the longitudinal axis of the c e l l . Microtubule bundles follow closely the curvature of the cell's surface. This i s noticeable at points where the vegetative filament branches (Fig. 17), and in the apical dome of the c e l l where they form a loose network (Fig. 19). Branching or reticulation of the microtubule network i s not seen in other areas of the c e l l , and the bundles have no obvious physical association with the majority of the organelles . However, an interaction between microtubule bundles and chloroplasts seems to exist, as indicated by their close association with extruded cytoplasm preparations (Fig. 23). There are no obvious focal points, similar to those observed for microfilaments, from which the microtubule bundles seem to. originate. Successive optical sections taken with the confocal laser scanning microscope show that microtubule bundles occur throughout the f u l l width of the cytoplasm in vegetative filaments (Fig. 21). This is confirmed by Figure 20 that shows a cross-sectional scan of microtubule bundle fluorescence. Figure 22 shows a transverse section with punctate B-tubulin fluorescence (presumably microtubule bundle ends) occupying the f u l l width of the cytoplasm of the vegetative filament to a depth (of up to about 15 urn) close to the vacuole. High resolution scanning electron microscopy studies Figure 24 shows a low magnification high resolution scanning electron micrograph of a transverse fractured face of a filament of Vaucheria longicaulis var. macounii. Chloroplasts occupy most of the vegetative filament cytoplasm. These organelles and their internal membranes seem to be well preserved. Two types of fibrous structures have been observed in such preparations. In Figure 24 (arrowhead) a f i b r i l l a r network can be discerned. At higher magnification, this f i b r i l l a r network appears as an irregular web of fibrous elements connecting different organelles (Fig. 25). These fibrous elements measure approximately 50 nm in diameter. Spherical bodies, measuring 50-100 nm in diameter, are frequently found in clusters in association with the fibrous elements of this network. Figure 26 shows another type of cable-like fibrous strand in the cytoplasm of a vegetative filament of Vaucheria. In this case, the individual elements are unbranched, straight and of determinate length. Figure 27 shows a higher magnification of these structures. The thinner elements measure approximately 50 nm in diameter and the wider ones 100 nm. Their interaction with organelles seems to be better defined than in the case of the web-like elements. Discussion - Part 1 The cytoskeleton of Vaucheria species has previously been studied by both di f f e r e n t i a l interference contrast (DIG) optics and electron microscope studies (Ott and Brown 1972, Blatt and Briggs 1980, Oliveira and Fitch 1988, Ott 1992, Tornbom and Oliveira, unpublished). Images of arrays of parallel fibrous cables observed in close association with organelles by differential interference contrast microscopy are similar to those seen in this study. Ott and Brown (1972) and Oliveira and Fitch (1988) showed using transmission electron microscopy, that microtubules are organized in bundles of 10-20 units, either in close contact with nuclei or the surface membrane of the c e l l . Microfilaments are not easily visualized by transmission electron microscopy, apparently due to the disruptive effects of aldehyde fixatives (Ott and Brown 1974, Blatt et a l . 1980, Doonan et a l . 1988, and our own observations). Kengen and de Graaf (1990) suggested that the s t a b i l i t y of actin during fixation may be dependent upon i t s interaction with other cytoskeleton proteins (e.g. microtubules or actin binding proteins), and this is worth bearing in mind when considering the organization and distribution of Vaucheria actin. Overall, the studies have been rather fragmented and no clear understanding of the dynamics of the organization of the cytoskeleton exists in this genus. In this study SDS-PAGE and immunoblots confirm that two of the major cytoskeletal proteins, actin and tubulin, are present in this alga. Commercially obtained monoclonal antibodies, raised against mammalian forms of the proteins cross-react with them demonstrating conserved epitopes. The actin molecule i s a highly conserved one (La Claire 1991), and in Vaucheria i t appears to be of similar molecular weight to actins from animal sources (43 kDa) and plant c e l l s such as Nitella (46 kDa, Allen and Allen 1978) and tomato (42 - 45 kDa, Seagull 1989). The upper band cross-reacting with the anti-B tubulin i s of a similar molecular weight to bovine brain tubulin, while the lower band of approximately 46.5 Mr i s probably a proteolysis breakdown product of B-tubulin. The same antibodies have also been used to demonstrate the presence of actin in Chara (Williamson et a l . 1987), Pisum (Abe and Davies 1991), pollen tubes (Tang et a l . 1989) and the fungus Phytophthera (Temperli et a l . 1990), and tubulin in Allium (Gubler 1989) and Phytophthera (Temperli et a l . 1990) - These antibodies are therefore suitable for identification and localization of actin and tubulin containing structures by immunofluorescence in Vaucheria filaments. There i s now a substantial body of evidence for the existence of an actomyosin system in plant and algal c e l l s , especially in association with cytoplasmic movement and wound healing (Williamson 1986, Williamson et a l . 1987, Kohno and Shimmen 1988, La Claire 1991). Our results indicate that myosin i s also present in Vaucheria. However, this i s known to be a very labile protein and not as highly conserved as actin (Parke et a l . 1986, La Claire 1991). Only a very faint band of approximate molecular weight 116 Mr showed a cross-reaction with anti-Dictyostelium myosin heavy chain polyclonal antibody. This antibody was shown to cross react with proteins, probably including myosin, in the alga Ernodesmis (La Claire 1991). The 55 Mr band, which cross-reacts with the commercial monoclonal anti-^pan' myosin, i s li k e l y a proteolysis breakdown product. This anti-myosin has been shown to cross-react with higher plant myosin (Parke et a l . 1986). At present, the absence of a recognizable myosin cross-reaction renders these antibodies unsuitable for myosin identification and localization by immunofluorescence in Vaucheria filaments. Phalloidin and immunofluorescence-labelling give clear images of actin and tubulin distribution in vegetative filaments of Vaucheria. respectively. Actin immunofluorescence did not give satisfactory results, perhaps due to d i f f i c u l t y of penetration of the large Amersham IgM monoclonal anti-actin antibody (La Claire 1989). Cells fixed with aldehyde after phalloidin stabilization show the distribution of the F-actin arrays to be similar to that of c e l l s permeabilized with low concentration DMSO, known to improve fine staining of F-actin arrays (see materials and methods for references). It i s also important to note that in the present study, the distribution and organization of fluorescent microfilament bundles i s similar to that of fibrous structures seen by differential interference contrast microscopy of living specimens. Ott (1992) describes the existence of a membranous ER-like compartment (motility associated reticulum or MAR), which he proposes to be equivalent to the Die cytoplasmic tracks and to participate in cytoplasm and organelle motility. The M^AR' elements were shown to .be associated with a diffuse actin-like network. Our observations tend to support Ott's (1992) conclusions that the M^AR' i s co-localized with microfilaments in vegetative filaments of Vaucheria. However, bearing in mind that Blatt et a l . (1980) presented good evidence for the presence of F-actin bundles of similar dimensions to those seen in this study, and that Ott (1992) himself proposes that microfilaments may form bundles during motility, our results showing the occurrence of microfilament bundles in Vaucheria more lik e l y represents the true configuration (Sonobe and Shibaoka 1989). The system of parallel actin cables running longitudinally to the cylindrical body of the vegetative filament in Vaucheria is similar in organization to that found in other elongated plant and algal c e l l s exhibiting cytoplasmic streaming (Palevitz and Hepler 1975, Parthasarathy et a l . 1985, Menzel and Schliwa 1986a, Tewinkel et a l . 1989, Jackson and Heath 1990). The microfilament cables of Vaucheria are approximately 0.37 jum in diameter, similar to estimates obtained from other plant and algal cells (Kersey and Wessels 1976, Blatt et a l . 1980, Parthasarathy and Pesacreta 1980). Estimates of approximately 60 individual microfilaments per cable in Vaucheria are also similar to those of Chara actin cables (Staiger and Schliwa 1987). High resolution scanning electron micrographs (HRSEMs) show an irregular web-like fibrous array similar to the fine, reticulate fluorescence of F-actin, and with dimensions that could correspond to bundles of several microfilaments. However, estimates of numbers of microfilaments per bundle carried out with fluorescence observations may be different due to the ^ cone' effect of fluorescent light emissions that increases the overall diameter of the cable-like structures. In addition, metal coating in HRSEM can add several nanometers to the dimensions of structures visualized by this technique (Ip and Fischman 1979). The microfilament array of Vaucheria seems to be composed of two sub-sets of cables: one made up of f a i r l y straight elements, oriented parallel to the longitudinal axis of the filament and adjacent to the plasma membrane, and beneath i t a reticulate set in close association with organelles, particularly chloroplasts. This observation i s supported by confocal scan images which reveal a more reticulate F-actin array, at a greater depth in the cytoplasm than the axially aligned microfilament cables. Two similar populations of actin microfilaments were observed in fern protonemata (Kadota and Wada 1989). Wasteneys and Williamson (1991) also observed distinct actin microfilaments in Nitella. with sub-cortical cables associated with chloroplasts and encircling nuclei which rotate in this species. One possible interpretation i s that the straighter microfilament bundles provide guiding tracks, and the reticulate ones, in association with organelles, enable them to move along those tracks. This could be specially true for chloroplasts, that show a very close association with the reticulate microfilament bundles located beneath the straighter set. In Vaucheria chloroplasts frequently reverse their direction of travel, implying the existence of microfilaments of different polarities within the array (Tornbom and Oliveira, unpublished). The existence of two sets of microfilaments could then provide a system for the regulation of complex organelle movement. Large coenocytic algae such as Vaucheria may require the a b i l i t y to accomplish sudden directional changes, as in response to wounding (Tornbom and Oliveira, unpublished). Large crystalline-like arrays of microfilament bundles occasionally seen in Vaucheria filaments seem to arise from, the aggregation of actin as a result of osmotic stress (Suzuki et a l . 1989). Similar structures, visualized using phalloidin staining, have been documented by Heslop-Harrison et a l . (1986), who postulated they were the result of F-actin storage in linear deposits. Pierson et a l . (1986) reported actin f o c i in pollen tubes which are ^dense and star shaped', and suggested a possible role in actin organization although acknowledging that they could be stress fibres. The crystalline-like actin aggregates in Vaucheria are distinct from the microfilament foci seen throughout the microfilament array. There are a few other reports of actin foci in the literature, but these show l i t t l e similarity to those observed in Vaucheria (Menzel 1987, Quader and Schnepf 1989). The role of foci in F-actin organization w i l l be considered in Part 2 of this study. The organization of F-actin in the apical dome of the t i p region of Vaucheria i s characterized by dense reticulation associated with intense pools of fluorescence, possibly f o c i . A similar organization observed at the apex in other tip-growing cel l s i s consistent with a role in exocytosis of vesicles carrying materials needed for the expansion of the c e l l wall (Schnepf 1986, Segawa and Yamashina 1989, Steer and Steer 1989). Microtubule bundles are longitudinally oriented to the cylindrical body of Vaucheria filaments. The short, straight microtubule bundles visualized in this study are consistent with Ott's (1992) description of a ^microtubular probe', attached to the anterior end of each nucleus and apparently involved in i t s motility. The close association of nuclei and microtubule bundles in Vaucheria (Ott and Brown 1972) and their response to the microtubule depolymerizing agent Oryzalin (Tornbom and Oliveira, unpublished) give strong evidence supporting a role for microtubules in nuclear motility. Vaucheria i s a tip-growing c e l l (Kataoka 1975) and similar organizations of microtubule arrays are seen in other cells exhibiting t i p growth (Derksen et a l . 1985, Doonan et a l . 1988, Temperli et a l . 1990). Therefore, the common features of microtubule arrays of tip-growing cells imply integral functions related to the formation and maintenance of a polarized c e l l structure (Lloyd 1984, Schnepf 1986). Oliveira and Fitch (1988) observed the constant presence of a group of up to 20 nuclei behind the apical dome of the t i p of the Vaucheria filament, while hundreds of other nuclei are distributed randomly throughout the remaining cytoplasm. These nuclei are associated with a loose network of microtubules present in the t i p region. In this manner they may also participate in the establishment of conditions preserving the integrity of the apical region (Doonan et a l . 1988). Indeed, nuclei have been shown to influence the site of t i p i n i t i a t i o n and to be transported equidistant to growing tips in a number of tip-growing systems (see Schnepf 1986 for review), although a direct role of nuclei in the establishment of polarity could not be demonstrated so far. There are no obvious focal regions for microtubule organization in Vaucheria. However, bright B-tubulin fluorescence spots observed in association with nuclei are consistent with the possibility that they may represent nucleation or polymerization sites (Part 2). Observations of depolymerization of tubulin after cold treatment, revealing a pool of fluorescence at the end of each shortened microtubule bundle are also consistent with this interpretation. Further support comes from studies showing similar pools of fluorescence in repolymerizing microtubule bundles experiments (Part 2). L i t t l e co-distribution of microtubule and microfilament arrays i s seen in Vaucheria. probably due to the marked difference in numbers of elements of the two arrays. However, the effect of both Cytochalasins and Oryzalin on organelle behaviour and cytoplasmic streaming suggests that both are involved in c e l l motility and intracellular organization (Tornbom and Oliveira, unpublished). Cross-bridging molecules such as MAP-2 could link the two systems as suggested in Acetabularia (Menzel and Elsner-Menzel 1989) and microtubule bundles could also serve as a template for orientation and a stabilizer of microfilament cables and the acto-myosin motor system (Hepler and Palevitz 1974, Menzel and Schliwa 1986a, La Claire 1987). Results - Part 2 Untreated c e l l s Figures 2.1a and b are differential interference contrast (DIG) micrographs, taken at an interval of approximately one minute apart, of a portion of a vegetative filament of Vaucheria lonqicaulis var. macounii. The cytoplasm i s organized into a network of cable-like strands (tracks) which seem to radiate from focal regions (arrows). Cytoplasmic tracks are not static within the c e l l ; hence, their reorganization from focal regions i s in a constant state of flux. Observed with fluorescein-phalloidin (Ph-FITC) staining, these regions appear as bright fluorescent areas from which F-actin cables seem to radiate (Fig. 2.2). The size of these foci i s quite variable and they can measure from 1 to 6 microns in diameter. In comparison, f o c i - l i k e structures visi b l e in DIC microscopy measure approximately 2 to 5 |xm. No clear relationship seems to exist between organelles and the foci, although the association of organelles with cytoplasmic tracks i s well defined when observed by DIC microscopy (Part l ) . No foci similar to those observed for microfilaments have been identified in association with the microtubule bundle array of the vegetative Vaucheria filament (Fig. 2.3). However, fixed ce l l s , sectioned with a freeze-stage microtome, often produce oblique cut ends which reveal an image of the cytoplasm viewed from either the plasma membrane or the tonoplast sides. These reveal punctate fluorescence when viewed from the tonoplast side (Fig. 2.4a) and filamentous fluorescence when viewed from the plasma membrane side (Fig. 2.4b). Given the fact that the predominant orientation of the microtubule bundles i s parallel to the c e l l surface and longitudinal axis of the vegetative filament (Fig. 2.3), these punctate structures possibly represent aggregates of B-tubulin sub-units which could function as microtubule-associated focal regions. F-actin foci In order to determine whether foci are related to the assembly and organization of the microfilament network, the cel l s were treated with Cytochalasins (F-actin inhibitors). Most cytoplasmic streaming ceases after 5 to 10 minutes of incubation with 10 jug/ml Cytochalasin D. Tornbom and Oliveira (unpublished) found that a much higher concentration (200 /xg/ml) of Cytochalasin B was required to stop cytoplasmic streaming almost immediately. Streaming i s restored 15 to 20 minutes after transfer to Cytochalasin-free growth medium. No change of c e l l shape i s observed upon treatment with Cytochalasins. Fluorescence microscopy reveals that treatment of the cells with Cytochalasins B and D results in the breakdown of the microfilament bundle array. The extent of the breakdown depends on the concentration and duration of the treatments. Cytochalasin D has a stronger effect, with very l i t t l e F-actin fluorescence observed when applied at 100 /lig/ml (Fig. 2.5). Cytochalasin B requires a much higher concentration (200 /xg/ml) to achieve a visib l e breakdown of the F-actin array (Fig. 2.6). When applied at 10 /xg/ml for approximately 10 minutes, Cytochalasin D was observed to cause breakdown of the microfilament array and hence i t was routinely used at this concentration in a l l the subsequent studies. The pattern of fluorescence observed after treatment with Cytochalasin D takes several forms, which may represent different stages of the breakdown of the F-actin array. These may result from the fact that individual c e l l s may react to drugs differently so as to produce different rates of depolymerization (Cleary and Hardham 1988) . The f i r s t indication of a disruption of the microfilament array i s the appearance of specks of brighter fluorescence amidst the axially aligned cables (Fig. 2.7, arrows). The F-actin array is now more reticulate and disorganized when compared to controls (Fig. 2.8). Intense, reticulate F-actin fluorescence becomes particularly apparent adjacent to cross walls formed during the sealing off of old wound sites (Fig. 2.9). Foci are visi b l e amidst the remaining F-actin cables (arrowheads). These focal regions are similar to those observed in control images (Fig. 2.2) and compare well with similar DIC structures (Fig. 2.1). Figure 2.10 shows another aspect of F-actin foci situated intermittently along microfilament cables (arrowheads). This pattern of F-actin depolymerization in close association with focal regions, i s seen to extend throughout most of the length of Cytochalasin-treated vegetative filament (Fig. 2.11, arrow), and into the t i p region (Fig. 2.12). Figure 2.13 shows a region of a c e l l with l i t t l e reticulate fluorescence, but with several F-actin cables that appear to be merging into a larger extremely bright structure. These and other similar structures are seen to radiate from a bright fluorescent spot (arrow). Extensive depolymerization and disruption of F-actin i s seen in Figure 2.14, where there are large areas of the c e l l without any F-actin, except for those areas associated with f o c i . Figure 2.15 shows a more advanced stage of F-actin depolymerization with only very compact and brightly fluorescent foci remaining. The microfilament array becomes highly disorganized after cold temperature treatment (< 0 °C), with the bundles of microfilaments losing their uniform orientation parallel to the longitudinal axis of the c e l l and displaying a more irregular configuration. The disorganized remnants of F-actin filaments are seen to be closely associated with numerous f o c i - l i k e regions (Fig. 2.16, arrows). The microtubule depolymerizing herbicide Oryzalin has no v i s i b l e effect on the microfilament array of the vegetative filament, as seen by Ph-FITC labelling (not shown). Recovery of Vaucheria cells after exposure to inhibitors was measured by the restoration of cytoplasmic streaming. Fluorescence images reveal microfilament arrays in the process of reassembly. The elements of this array are better defined and clearly associated with focal regions (Fig. 2.17, arrows). Similar F-actin arrays containing foci are observed after re-warming of cold-treated ce l l s , again suggesting that the new arrays are in the process of reassembly (not shown). Microtubule-associated foci Cells were treated with cold temperature, the herbicide Oryzalin, and the stabilizing agent taxol, to investigate the organization of the microtubule array and i t s possible relationship to the bright fluorescent B-tubulin spots observed amongst the microtubule bundle array (Fig. 2.4). The effects observed after cold and Oryzalin treatments are similar. The results reveal that Vaucheria c e l l s have cold-tolerant microtubules. Depolymerization was never observed unless c e l l s were treated with temperatures below 0 °C, and an apparently normal microtubule array was sometimes s t i l l present after exposure to temperatures as low as -3 °C (Fig. 2.18). Cytoplasmic streaming ceased at temperatures below 0 °C, and after treatment with 10 /nM Oryzalin. It was restored upon rewarming of the medium or upon placement of vegetative filaments in Oryzalin-free growth medium. Recovery occurred within a period of 30 minutes to one hour. Incomplete depolymerization of the microtubule bundles by these treatments reveal intermediate stages of breakdown. The f i r s t indication of depolymerization is marked by the appearance of fragmented microtubule bundles (Fig. 2.23). In this process, bright spots of fluorescence become visib l e at one end of the microtubule bundle (Fig. 2.. 25, arrow). Other effects of incomplete depolymerization include disorientation of short microtubule bundles so that they l i e at an angle to the longitudinal axis of the c e l l . Some bright spots of fluorescence are observed amidst the increasingly disorganized array (Fig. 2.24 arrows). Complete depolymerization of microtubules either by cold temperature or Oryzalin treatments result in the breakdown of microtubule cables into a series of localized bright spots of fluorescence. These sometimes occur in rows, as though the whole cable has been fragmented into multiple pools of B-tubulin (fig. 2.19). Figure 2.20 shows the t i p region of a vegetative filament after complete depolymerization of microtubules by cold treatment. Once more, only punctate fluorescence i s observed. In some cases, the location of these pools of tubulin correlates well with the positions of nuclei (as visualized using DAPI staining) (Figs. 2.21 and 2.22). Upon closer inspection, the tubulin staining sometimes appears to be bipolar in relation to each tear-shaped nucleus. Nuclear autofluorescence was never observed. The microtubule array remains v i s i b l e after incubation in the stabilizing agent taxol, but some disorganization is apparent. This appears in the form of wavy and brightly fluorescent spot-terminated cables (Fig. 2.26). Taxol did not stabilize the microtubules against cold depolymerization (data not shown). Cells exhibiting cytoplasmic movement after re-warming revealed microtubule arrays in different stages of apparent repolymerization and reorganization. The f i r s t indication of repolymerization i s evident in the formation of large areas of bright B-tubulin fluorescence in association with bright f o c i -like spots (Fig. 2.27, arrowhead). Curved, short microtubules are also observed in contact with bright spots of fluorescence at a more advanced stage of tubulin polymerization (Fig. 2.28). Figures 2.29 and 2.30 show an apparent progressive formation of microtubule bundles oriented parallel to the longitudinal axis of the vegetative filament. Cytochalasins show no effect on the structure or distribution of the microtubule array (not shown). Discussion - Part 2 Part 1 reported the existence of extensive arrays of axially aligned microfilament cables and microtubule bundles in Vaucheria lonqicaulis var. macounii. Part 2 investigates how the organization of this large and complex cytoskeleton system might be accomplished, and the possible role(s) of focal regions as organizing centers. Cytochalasin D causes a step-like breakdown of the microfilament array of Vaucheria. This i s dependent upon the concentration and the duration of the treatments (Williamson 1978, Schliwa 1982, Yahara et a l . 1982, Tang et a l . 1989, Murali Krishna Rao et a l . 1992) and i t provides a useful tool to analyse the organization of the microfilament array. A consistent feature of the disorganization of the microfilament array is that i t always occurs in close association with regions of the F-actin network called f o c i . These observations are consistent with those of Blatt et a l . (1980) who observed reticulation and branching of cortical fibers by differential interference contrast microscopy after application of Cytochalasin B. Yumura and Kitanishi-Yumura (1990) observed similar structures in Dictyostelium and proposed them to be organizing centers of F-actin. Formation of F-actin aggregates in plant c e l l s i s common under the influence of Cytochalasin D (Parthasarathy 1985, Menzel and Schliwa 1986b, Palevitz 1988). Differential interference contrast observations of livi n g vegetative filaments show that the cytoplasmic track network seems to originate from highly dynamic focal regions which appear to have a similar distribution to the F-actin foci (Part 1). Therefore the F-actin focal regions are not l i k e l y to be an artifact of the methods of preparation. Foci may be formed by contraction of F-actin. This i s proposed to be mediated by actin binding proteins located within the focal regions (Schliwa 1982, Yahara et a l . 1982, Yumura and Kitanishi-Yumura 1990). It i s interesting to notice in relation to this interpretation that images show that as depolymerization progresses, foci seem to increase in size concurrent with the alteration and eventual disappearance of the F-actin cables. The probable presence of myosin, in association with the microfilament array (Part 1) may allow for a similar phenomenon to occur in Vaucheria. Repolymerization studies further support the existence of F-actin foci in association with F-actin cables and suggest the possibility that they act as polymerization centers. Yumura and Kitanishi-Yumura (1990) showed that in Dictvostelium foci are located on the plasma membrane. In Vaucheria vegetative filaments, the location of the whole F-actin array, including foci, in the cortical cytoplasm suggest that the plasma membrane may have an organizational function (Traas 1990). A possible model for this type of organizational structure could include actin binding integral membrane proteins similair to those known to occur in animal cells, such as ponticulin (Wuestehube and Luna 1987) and vinculin (Jockusch and Isenberg 1981). These molecular interactions could act to transmit signals via the plasma membrane to the closely associated cytoskeleton (Isenberg 1991). This would enable the c e l l to co-ordinate the distribution of microfilament bundles in the coenocytic vegetative filament of Vaucheria. The existence of multiple foci distributed throughout the microfilament array in untreated c e l l s of Vaucheria is consistent with this interpretation. These signals could participate in the organization of F-actin into two distinct, functionally different sub-sets of microfilament cables (Part 1). However, no preferential localization of foci between the two sub-sets of F-actin has been observed. It i s possible then that polymerization at the foci may not distinguish between different sub-sets of microfilaments. Foci and extensive arrays of F-actin are observed close to cross-walls formed in response to wounding. Under these circumstances, no differentiation between the two sub-sets of F-actin cables can be seen, suggesting that this may be occurring after polymerization. The effects of Oryzalin and cold temperature on cytoplasmic movement in Vaucheria are reversible. These inhibitors depolymerize microtubule bundles into a series of localized spots of tubulin, similar in appearance to the punctate fluorescence observed in untreated c e l l s . This effect i s also observed in other plant c e l l s where i t i s interpreted as indicative of microtubule depolymerization (Menzel and Schliwa 1986b, Cleary and Hardham 1988, Wacker et a l . 1988, Wasteneys and Williamson 1989, Akashi et a l . 1990, Schwuchow et a l . 1990, Astrom et a l . 1991) . Microtubule depolymerization in response to cold treatment i s particularly pertinent for this investigation because of i t s relevance to the natural habitat of Vaucheria. where low temperatures within the range used in this study are frequently experienced during the winter. Transmission electron microscopy of a variety of species of Vaucheria suggests a close relationship between microtubule bundles and the anterior end of each nucleus, including a pair of centrioles proposed to act as a microtubule organizing centre (MTOC) (Ott and Brown 1972, Ott 1992) . Some of the brightly fluorescent spots (foci) seen after depolymerization appear to be co-localized with nuclei. These are also observed in repolymerization studies, which indicates nucleation of tubulin at these sites. Nuclear-related, MTOC anchored, and ^capped' microtubules are cold-stable (Doonan et a l . 1988, McBeath and Fujiwara 1990), and this i s consistent with our observations of the extreme tolerance of Vaucheria microtubules to cold temperatures. Taxol causes the appearance of microtubule bundles in association with bright spots (foci). Similar effects of taxol were suggested to be due to reorganization of microtubules through increased bundling and aggregation of tubulin (Kuss-Wymer and Cyr 1992), and establishment of microtubule cross-links (Melan 1990). These observations further supports the interpretation that they are involved with the polymerization and organization of tubulin. Polymerization may be occurring at the d i s t a l (plus) end (McBeath and Fujiwara 1990), i f the microtubule bundle is anchored at the centrioles (MTOC). In Vaucheria. nuclei do not divide synchronously (Ott and Brown 1972). Therefore the nucleus associated tubulin fluorescence i s not always observed. Microtubule-associated fluorescent spots in Vaucheria which do not co-localize with nuclei may represent other nucleation sites, such as those found in regions adjacent to the plasma membrane (Falconer et a l . 1988). This i s supported by Ott's (1992) observations that not a l l microtubule bundles in Vaucheria seem to be attached to nuclei. In this case i t i s possible that microtubule bundles are organized by a nucleus-associated MTOC that detaches from i t at a certain point during their growth (McBeath and Fujiwara 1990). The existence of ^ microtubule probes' recently described by Ott (1992) in Vaucheria longicaulis may be the result of such a mechanism. Our repolymerization studies did not reveal radial growth of microtubules from focal points (Lloyd 1987). Instead, longitudinally oriented microtubule bundles with unidirectional growth are seen to be associated with spot-like f o c i . This organizational pattern may be the result of the presence of MAPs which cause increased bundling of polymerizing microtubules (Wasteneys and Williamson 1989), and perhaps a ^pre-programmed' pattern of par a l l e l growth at the MTOC (Brown et a l . 1982, Kalnins 1992). This pattern of growth of microtubule bundles i s consistent with the role they and associated nuclei are thought to play in the polarized growth of Vaucheria vegetative filaments (Kataoka 1975, Part 1). The lack of effect of Cytochalasins, on the organization of the microtubule array, and Oryzalin, on the microfilament array, is interesting when considering the proposed interaction between the two cytoskeleton systems in organelle movement (Ott 1992, Tornbom and Oliveira, unpublished. Part 1). Cold treatment did, however, depolymerize the microfilament cables to reveal f o c i . These observations suggest that interaction between the two arrays may be mediated by proteins such as MAPs (Pollard et a l . 1984, Menzel and Elsner-Menzel 1989) which allow the two systems to function in tandem, whilst enabling them to maintain specialized roles (Ott 1992, Tornbom and Oliveira, unpublished). These specializations are reflected in the differences in their organization from distinct nucleating centers. A nucleus and i t s associated MTOC seem to organize the microtubule bundle as an independent unit with growth from the distal end of the microtubule, whereas F-actin cables seem to be organized at the proximal end from centers distributed throughout the cytoplasm (Dustin 1984). Conclusion This study has identified two cytoskeleton components, microtubules and microfilaments, and described their overall organization in vegetative filaments of Vaucheria lonqicaulis var. macounii. The extensive array of longitudinally oriented microfilaments, and the less dense array of microtubules undoubtedly play important roles in the polarized organization of this coenocytic alga which exhibits vigourous cytoplasmic streaming. There i s also evidence for the existence of the mechanomotor myosin, although not as conclusive as for the other two components. Experimental manipulation of these cytoskeleton elements, through disruption (depolymerization) and recovery (repolymerization), gives some insight into how control over the arrays of cytoskeletal elements may be accomplished. This seems to occur through organizing centers with different properties and distributions for microfilaments and microtubules. Microfilaments may be organized by foci localized at sites adjacent to the plasma membrane and distributed throughout the filament, whereas microtubule bundles are thought to be organized at centers associated with the centrioles and nuclei. Nuclei-free organization centers are also observed and thought to result from detachment of microtubules from their original organizing center. An obvious direction in which to continue this study is to unequivocally identify myosin, and to probe for other proteins such as MAPS and intermediate filament related proteins using SDS-PAGE and immunoblotting. Localization of these proteins using immunofluorescence confocal microscopy w i l l add further depth of knowledge to the organization and functioning of the cytoskeleton. Treatment of vegetative filaments with N-ethylmaleimide, a dynein and myosin poison, w i l l provide further evidence as to the presence of molecular motors and the role(s) they play in this organism. Immunofluorescence studies of cellular extracts w i l l add to a better identification of the relationship of different organelles with both F-actin and microtubule bundles. In addition, cellular extracts, as used for other algal systems (Kohno et a l . 1990), may provide a useful tool for the study of the role different cytoskeleton molecules play in organelle transport and cellular organization. Microscopical work involving labelling of F-actin with heavy meromyosin for TEM w i l l provide information on the polarity of the microfilaments, which is important when considering the motility mechanism. Immunoelectron microscopy (TEM and HRSEM) can also be used to locate some of the proteins discussed in this study and their relationship to the putative organizing centers (foci), while providing at the same time knowledge about their ultrastructural organization. FIGURES - PART Fig. 1 SDS-PAGE gels and immunoblots of Vaucheria homogenate proteins A. 12% gel of Vaucheria proteins (a), anti-actin immunoblot of lane a (b), negative control (nitrocellulose treated only with secondary antibody) (c) B. 12% gel of Vaucheria proteins (d), bovine brain tubulin standard (e), anti-tubulin immunoblot of lane d (f) , anti-tubulin immunoblot of lane e (g) G. 12% gel of Vaucheria proteins - note faint band at approximately 200 kDa (arrowhead) (h), anti-myosin (Amersham) immunoblot of lane h (i) D. 7.5% gel of Vaucheria proteins - note band at approximately 200 kDa (arrowhead) (j ) , anti-Dictyostelium myosin immunoblot of lane j - note faint skewed band at approximately 110 kDa (arrowhead) (k), anti-Dictyostelium myosin immunoblot of myosin standard (1), negative control (nitrocellulose treated only with secondary antibody) (m) Figs. 2 to 7 Bars = 10 jum Figs. 2 & 3 Differential interference contrast microscopy of a Vaucheria filament. Figure 2 shows an axially aligned array of fibrous structures distributed throughout the cytoplasm, in close association with c e l l organelles such as chloroplasts (arrow) and vesicles (arrowheads). Figure 3 shows a different region of a Vaucheria filament, demonstrating the relationship between the fibrous structures and chloroplasts as well as nuclei (arrowheads) Fig. 4 F-actin microfilament array visualized using fluorescein-labelled phalloidin (unfixed material). A dense network of parallel, axially aligned cables can be seen. Inset: Distribution of F-actin by focussing on the cortical region of the filament, showing that microfilament cables are present within approximately 2 fj.m of the cel l ' s surface Fig. 5 Microfilament array of an aldehyde-fixed c e l l showing branching of longitudinal cables in the cytoplasm (arrowhead) and around a chloroplast (c) (arrow) Fig. 6 Branching of microfilament cables in close proximity to organelles (arrowheads) in an unfixed filament stained with fluorescein-labelled phalloidin Fig. 7 Crystalline-like microfilament arrays in an osmotically stressed filament of Vaucheria Fig. 8 Fluorescein-labelled phalloidin staining shows the association of microfilament cables and chloroplasts (c) . Bar = 10 /Ltm Fig. 9 Microfilament cables appear to radiate from a focal area (arrow) in the apical region of a vegetative filament stained with fluorescein-labelled phalloidin. Bar = 10 /im. Inset: intense, fine F-actin fluorescence in the t i p region of a filament. Bar = 5 /xm Fig. 10 Microfilament cables appear to occur in two forms: straighter ones, parallel to the longitudinal axis of the c e l l and adjacent to the cel l ' s surface (arrows), and more reticulate ones in apparent association with organelles and located deeper in the cytoplasm (arrowheads). Bar = 10 r^o Fig. 11 Focal point of microfilament cables with many radiating from and becoming parallel to the longitudinal axis of the c e l l . Bar = 10 fjLm. Inset: focal point with fewer microfilament cables. Bar = 5 fim Fig. 12 Confocal scans (sequence l e f t to right) at intervals of 0.8 njR, showing the partitioning of the microfilament array throughout the cytoplasm of a Vaucheria filament labelled with fluorescein-labelled phalloidin. Bar = 50 nm Fig. 13 Confocal scans (sequence l e f t to right) at intervals of 1 fim (upper series, and upper to lower series) and 2 /xm (lower series). Fluorescein-labelled phalloidin staining of a vegetative filament shows longitudinally aligned microfilaments within the f i r s t 2 nm of the ce l l ' s surface, and reticulated regions of actin microfilaments at a depth of approximately 4 nm into the cytoplasm. Focal regions are also observed in a l l levels of optical sectioning. Bar = 50 n^ Fig. 14 Confocal cross-sectional scan of a fluorescein-labelled phalloidin stained vegetative filament, with graphical representation of fluorescence distribution within the cytoplasm. The highest density of fluorescence occurs in the cort i c a l region. Bar = 10 m^ Fig. 15 Confocal graphical representation of fluorescence distribution along a labelled longitudinal axis of a vegetative filament. A similar pattern of fluorescence distribution seems to occur throughout the filament. Bar = 10 m^ Figs. 16 to 19 Bars = 10 jum Fig. 16 Microtubule bundles are visualized using indirect immunofluorescence for B-tubulin. They show axial orientation and determinate length, and sparse distribution throughout the cytoplasm Fig. 17 Vaucheria c e l l showing microtubule bundles curving and crossing over into a branch (arrowheads) Fig. 18 Confocal composite image of 30 scans showing the overall distribution of microtubule bundles in the mid-section of a vegetative filament. Fig. 19 Microtubule bundles form a loose network in the apical dome of the t i p region of a filament Figs. 20 to 23 Bars = 10 nm, except Fig. 21 Bar = 50 /um Fig. 20 Cross-section confocal scan and graphical representation of B-tubulin fluorescence distribution. High fluorescence intensity i s not equally distributed throughout the cytoplasm. Fig. 21 Serial confocal scans at depth intervals of 3 jum ( f i r s t scan at a depth of 10 /ia into the cytoplasm) . Microtubule bundles show that they are present throughout most of the cytoplasm width. Fig. 22 Cross-section of a Vaucheria filament showing punctate microtubule fluorescence (arrowheads) deeper into the cytoplasm. Fig. 23 Extruded chloroplast (c) with closely associated microtubule bundles (arrowheads). Fig. 24 Fractured end of a Vaucheria c e l l as seen by HRSEM. Chloroplasts (c), c e l l wall (cw) and fibrous material (arrowhead) are visible. Bar = 5 Fig. 25 Higher magnification of fibrous material in Figure 24 showing i t s web-like appearance, attachment to organelles (arrowheads) and small vesicles (arrow). Bar = 5 /im Fig. 26 Fractured filament of Vaucheria showing cable-like structures within the cytoplasm. These structures are largely unbranched and run straight between different areas of the cytoplasm. Bar = 2.5 (xm Fig. 27 Higher magnification of Fig. 26 showing the association between an organelle (arrowhead) and thinner cables. Larger cables (arrow) appear to be made up of thinner cables. Bar = 150 nm FIGURES - PART Figs. 2.1 to 2.4 Bars = 10 jum Figs. 2.1a & b a. Differential interference microscopy micrograph showing a region of a vegetative filament adjacent to a cross-wall formed in response to wounding (arrow). A network of cytoplasmic tracks i s seen throughout the cytoplasm and these seem to radiate from focal areas (arrowheads). b. Same area of the c e l l one minute later, demonstrating that these are dynamic structures with anastamosing elements. Fig. 2.2 Reticulate F-actin array, visualized by fluorescein-labelled phalloidin, i s seen to be associated with numerous foci (arrowheads). These are observed adjacent to a cross-wall of a vegetative filament formed in response to wounding (arrow). Fig. 2.3 Microtubule array of a vegetative filament visualized using B-tubulin immunofluorescence. Figs. 2.4a & b a. Oblique cut end of a vegetative filament reveals filamentous and punctate B-tubulin fluorescence when focussing on the tonoplast side. b. Same filament shows only filamentous fluorescence when focussed from the plasma membrane side. Figs. 2.5 to 2.10 Bar == 10 fim, except Fig. 2.6 Bar = 5 /xm Fig. 2.5 Absence of F-actin fluorescence in a vegetative filament of Vaucheria after treatment with 100 /xg/ml Cytochalasin D. Some autofluorescence of chloroplasts, and small specks of fluorescein are v i s i b l e (arrowheads). Fig. 2.6 F-actin fluorescence of a vegetative filament after treatment with 200 /xg/ml Cytochalasin B. The F-actin array i s largely disorganized, with i t s remnants associated with foc i - l i k e structures (arrowheads). Fig. 2.7 The f i r s t indication of breakdown of the F-actin array upon treatment with 10 /xg/ml Cytochalasin D i s represented by bright specks of fluorescence (arrowheads) amidst the more irregular and reticulate F-actin cables. Fig. 2.8 F-actin array of an untreated vegetative filament. Note the overall axial alignment of the F-actin cables. Fig. 2.9 Region adjacent to a cross-wall (formed in response to wounding) after treatment with 10 /xg/ml Cytochalasin D. Large (arrow) and small (arrowhead) foc i of F-actin are visib l e amongst the remaining F-actin cables. Fig. 2.10 Foci of F-actin (arrowheads) are v i s i b l e along the length of F-actin cables after treatment with 10 /xg/ml Cytochalasin D. Figs. 2.11 to 2.16 Bar = 10 fim, except Fig. 2.12 Bar = 5 nm Fig. 2.11 Web-like array of F-actin with interspersed fo c i (arrow) i s seen throughout the mid-filament region of a vegetative filament after treatment with 10 /ig/ml Cytochalasin D. Fig. 2.12 Similar web-like array with interspersed foci i s seen to extend into the tip region of a vegetative filament. Fig. 2.13 Advanced stage of depolymerization of the F-actin array by Cytochalasin D. Axially aligned F-actin cables appear to merge into one, large brightly fluorescent structures that are seen converging into a central focal region (arrow). Fig. 2.14 Extensive depolymerization of F-actin. Large areas of the c e l l show no fluorescence, and only a few foci with short, disorganized F-actin cables remain. Fig. 2.15 F-actin depolymerization i s almost complete, except for a few intensely fluorescent compact f o c i . Fig. 2.16 Disorganization of the F-actin array of a vegetative filament after cold (< -2 °C) treatment. Foci appear amidst the disrupted F-actin cables (arrows). Figs. 2.17 to 2.22 Bar = 10 /xm, except Fig. 2.18 Bar = 5 /xm Fig. 2.17 Recovery from Cytochalasin D treatment. Bright fluorescent foci are visible (arrows), amidst partially reorganized F-actin cables. Fig. 2.18 Microtubule array present in a vegetative filament of Vaucheria after exposure to sub-zero temperatures (-2 °C). Fig. 2.19 Complete depolymerization of the microtubule array in a vegetative filament, after exposure to cold temperature (-2 °C). Fluorescence is vi s i b l e as a series of bright spots (arrowheads), often in linear alignment. Fig. 2.20 Complete depolymerizatin of the microtubule array in the tip region of a filament after exposure to cold temperature (-2 °C). Only bright spots of fluorescence are visible. Figs. 2.21 & 2.22 Demonstration of the co-localization of B-tubulin spots and nuclei (compare Figs. 2.21 and 2.22). Figure 2.21 shows punctate B-tubulin fluorescence after cold treatment (-2 °C). Figure 2.22 shows nuclei visualized by DAPI staining. The B-tubulin fluorescence displays a bipolar distribution with respect to the nuclei (arrowheads). Figs. 2.23 to 2.26 Bars = 10 /iia Fig. 2.23 F i r s t indication of depolymerization i s evident by the fragmentation of microtubule bundles. Fig. 2.24 Disorganization of microtubule bundles is accompanied by the appearance of bright fluorescent spots at their ends (arrows). Fig. 2.25 Bright spot of fluorescence at the end of a microtubule bundle (arrow). Fig. 2.2 6 A high degree of disorganization of microtubule bundles characterizes the treatment with taxol. 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