Tail chimeras of Dictyostelium myosin II support cytokinesis and other myosin II activities but not full development

Shi Shu1, Xiong Liu1, Carole A. Parent2, Taro Q. P. Uyeda3 and Edward D. Korn1,*

1 Laboratory of Cell Biology, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, MD 20892, USA
2 Laboratory of Cellular and Molecular Biology, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892, USA
3 Gene Function Research Center, Tsukuba Central #4, National Institute of Advanced Industrial Science and Technology (AIST), Higashi 1-1-1, Tsukuba, Ibaraki 305-8562, Japan

* Author for correspondence (e-mail: edk{at}nih.gov)

Accepted 21 August 2002


    Summary
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 Summary
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 Materials and Methods
 Results
 Discussion
 References
 
Dictyostelium lacking myosin II cannot grow in suspension culture, develop beyond the mound stage or cap concanavalin A receptors and chemotaxis is impaired. Recently, we showed that the actin-activated MgATPase activity of myosin chimeras in which the tail domain of Dictyostelium myosin II heavy chain is replaced by the tail domain of either Acanthamoeba or chicken smooth muscle myosin II is unregulated and about 20 times higher than wild-type myosin. The Acanthamoeba chimera forms short bipolar filaments similar to, but shorter than, filaments of Dictyostelium myosin and the smooth muscle chimera forms much larger side-polar filaments. We now find that the Acanthamoeba chimera expressed in myosin null cells localizes to the periphery of vegetative amoeba similarly to wild-type myosin but the smooth muscle chimera is heavily concentrated in a single cortical patch. Despite their different tail sequences and filament structures and different localization of the smooth muscle chimera in interphase cells, both chimeras support growth in suspension culture and concanavalin A capping and colocalize with the ConA cap but the Acanthamoeba chimera subsequently disperses more slowly than wild-type myosin and the smooth muscle chimera apparently not at all. Both chimeras also partially rescue chemotaxis. However, neither supports full development. Thus, neither regulation of myosin activity, nor regulation of myosin polymerization nor bipolar filaments is required for many functions of Dictyostelium myosin II and there may be no specific sequence required for localization of myosin to the cleavage furrow.

Key words: Myosin II, Cytokinesis, Chemotaxis, Development, ConA capping


    Introduction
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Class II myosins consist of two identical heavy chains and two pairs of light chains (Sellers, 1999Go). Because the cellular slime mold Dictyostelium discoideum contains only a single myosin II heavy chain gene (De Lozanne et al., 1985Go; Warrick et al., 1986Go), Dictyostelium myosin II heavy chain-null cells provide an excellent system for determining the biological roles of myosin II and the abilities of mutant myosin II heavy chains to substitute for wild-type heavy chain. In this way, it has been shown that Dictyostelium myosin II (DdMII) is required for cytokinesis and growth of vegetative amoeboid cells in suspension culture, for development and differentiation beyond the mound stage, the first developmental stage, (Knecht and Loomis, 1987Go; De Lozanne and Spudich, 1987Go; Manstein et al., 1989Go) and for capping of concanavalin A (ConA) receptors [(Pasternak et al., 1989bGo; Fukui et al., 1990Go) but see Aguado-Velasco and Bretscher, for a contrary view (Aguado-Velasco and Bretscher, 1997Go)]. DdMII heavy chain-null cells are motile and capable of cAMP-induced chemotaxis but both motility and chemotaxis in a spatial gradient of cAMP are impaired compared to wild-type cells (De Lozanne and Spudich, 1987Go; Wessels et al., 1988Go; Peters et al., 1988Go). Null cells can phagocytose bacteria but at the time the experiments described in this paper were carried out phagocytosis by null cells had not been quantified.

Cytokinesis in suspension culture, capping of ConA receptors and full development of Dictyostelium to fruiting bodies depend on both the actin-dependent MgATPase activity of DdMII and its ability to form filaments. For example, myosins with mutations in the motor domain of the heavy chain that eliminate ATPase activity but have no effect on polymerization competence (Yumura and Uyeda, 1997aGo) and polymerization-competent myosin rods devoid of the motor domain (Zang and Spudich, 1998Go) localize properly to the cleavage furrow but neither protein supports cytokinesis. On the other hand, myosin with mutations in the tail domain that prevent polymerization but have no effect on catalytic activity (Egelhoff et al., 1993Go) and catalytically active but polymerization incompetent heavy meromyosin (De Lozanne and Spudich, 1987Go; Fukui et al., 1990Go) fail to localize to the cleavage furrow and support neither cytokinesis, capping of ConA receptors nor full development. Recent experiments by Shu et al. indicate that no specific region of the rod other than the region within the light meromyosin domain that is required for filament formation is required for localization and function of myosin at the cleavage furrow (Shu et al., 1999Go). Therefore, myosin II filaments appear to be transported to the cleavage furrow by an unknown mechanism that is independent of the myosin motor II domain and of any specific sequence in most, and possibly all, of the tail domain.

Both the actin-activated MgATPase activity and filament formation of DdMII are regulated. Actin-dependent MgATPase activity of DdMII is increased about fivefold in vitro (Griffith et al., 1987Go; Uyeda et al., 1996Go; Liu et al., 1998Go) by phosphorylation of serine-13 on its regulatory light chain (Griffith et al., 1987Go; Ostrow et al., 1994Go); filament formation is blocked by phosphorylation of threonines-1823, 1833 and 2029 in the tail domain (Kuczmarski et al., 1987Go; Vailancourt et al., 1988Go; Luck-Vielmetter et al., 1990Go; Pasternak et al., 1989aGo; Liang et al., 1999Go); and changes in the level of phosphorylation of both light (Kuczmarski and Spudich, 1980Go; Berlot et al., 1985Go; Griffith et al., 1987Go; Berlot et al., 1987Go) and heavy (Malchow et al., 1981Go; Berlot et al., 1987Go) chains are associated with DdMII function in vivo. However, neither modulation of actin-dependent MgATPase activity nor regulation of filament formation is essential for DdMII function. For example, myosins in which regulatory light chain serine-13 has been mutated to alanine (Ostrow et al., 1994Go) or the light chain removed by deletion of the light chain-binding site on the heavy chain (Uyeda and Spudich, 1993Go; Yumura and Uyeda, 1997bGo), both of which have unregulated actindependent MgATPase in vitro (the former has 30% and the latter 240% of the activity of phosphorylated DdMII), support cytokinesis in suspension culture and full development to the fruiting body stage. Similarly, myosin in which the three phosphorylatable threonine residues in the tail are replaced by alanines (and which, as a consequence, over-polymerizes in vivo) supports cytokinesis in suspension culture, capping of ConA receptors and full development (Egelhoff et al., 1993Go), although growth is slower (Egelhoff et al., 1993Go) and motility is retarded (Yumura and Uyeda, 1997bGo) compared to cells expressing wild-type myosin. Also, truncated DdMII missing the C-terminal 34 kDa, which contains the phosphorylation sites that regulate filament formation, supports cytokinesis in suspension culture (Egelhoff et al., 1991aGo) and full development and capping of ConA receptors (O'Halloran and Spudich, 1990Go).

The results briefly summarized above show that myosin II function in Dictyostelium requires that the myosin be catalytically active and capable of forming filaments but that regulation of neither property is essential. Moreover, localization of myosin II to the cleavage furrow requires neither enzymatic activity, nor the motor domain nor any specific sequence in the tail other than, perhaps, within the region required for polymerization.

Recently, we (Liu et al., 2000Go) produced and characterized chimeric myosins consisting of the Dictyostelium myosin II heavy chain head and neck domains, with associated Dictyostelium light chains, fused to the tail domain of either chicken gizzard smooth muscle (Ch-Sm) or Acanthamoeba (Ch-Ac) myosin II (Fig. 1A). The unphosphorylated chimeras have about 20-fold higher actin-activated MgATPase than unphosphorylated DdMII in vitro resulting from a 10- to 15-fold increase in Vmax and a 2- to 7-fold decrease in KATPase (the concentration of F-actin required for half maximal activation). The activities of the chimeras are unaffected by light chain phosphorylation and are much greater than the activities of the other unregulated constructs of Dictyostelium myosin referred to earlier (Uyeda and Spudich, 1993Go; Ostrow et al., 1994Go; Liu et al., 1998Go).



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Fig. 1. Expression of Dd-Wt, Ch-Ac and Ch-Sm in HS1 null cells. (A) Schematic representation of Dd-Wt, Ch-Sm and Ch-Ac heavy chains (modified figure reproduced with permission from Liu et al., 2000Go). (B) Filaments of purified Dd-Wt, Ch-Sm and Ch-Ac myosins (reproduced with permission from Liu et al., 2000Go). (C) Coomassie-blue SDS-PAGE of total cell extracts of AX3 cells and HS1 cells expressing Dd-Wt, Ch-Ac or Ch-Sm. (D) Westerns of SDS-PAGE of total cell extracts. AX3, HS1 and Dd-Wt cells were blotted with anti-DdMII, Ch-Ac cells were blotted with anti-Acanthamoeba myosin II and Ch-Sm cells were blotted with anti-smooth muscle myosin II. Anti-Acanthamoeba myosin and anti-smooth muscle myosin did not react with HS1 cells (not shown).

 

Filaments of Ch-Ac resemble filaments of wild-type Acanthamoeba myosin II (Pollard, 1982Go), 200 nm long, bipolar filaments with a 90 nm central bare zone containing approximately 10-20 molecules, and are similar to but smaller than filaments of Dd-Wt (wild-type myosin expressed in null-cells), 450 nm long, bipolar filaments with a 150 nm central bare zone containing 25-50 molecules (Fig. 1B). The different lengths of the Ch-Ac and Dd-Wt filaments and their central bare zones reflect the differences in the lengths of the tail domains of the parental myosins and the number of molecules in the filaments. In marked contrast to the Dd-Wt and Ch-Ac filaments, filaments of Ch-Sm resemble filaments of smooth muscle myosin (Craig and Megerman, 1977), 700 nm long, side-polar filaments with no central bare zone and containing hundreds of molecules (Fig. 1B).

In the experiments described in this paper, we evaluated the ability of the two hyperactive and unregulated myosin chimeras to rescue the functional deficiencies of Dictyostelium myosin II-null cells. We were especially interested in the properties of Ch-Sm given the importance of filaments for myosin II localization and function and the very different structures, at least in vitro, of the large, side-polar Ch-Sm filaments and the small, bipolar Dd-Wt filaments.

We found that Ch-Ac localized similarly to Dd-Wt mostly to the periphery of vegetative amoeba whereas Ch-Sm was heavily concentrated in a single cortical patch. Despite their different localizations in interphase cells, different tail sequences and different filament structures, both chimeras supported cytokinesis and growth of DdMII-null cells in suspension culture. Both chimeric myosins supported capping of ConA receptors and both chimeras co-localized with the ConA cap (like wild-type myosin) but Ch-Ac redistributed much more slowly than Dd-Wt and wild-type myosin, and Ch-Sm apparently not at all, after capping was completed. Both chimeras partially rescued the reduced rates of motility and chemotaxis of DdMII-null cells and Ch-Ac rescued the mildly impaired phagocytic activity of DdMII-null cells. Neither chimera, however, supported full development.


    Materials and Methods
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Plasmid construction
All DNA manipulations were carried out using standard methods. The construction of chimeric myosin heavy chain consisting of Dictyostelium myosin II head and Acanthamoeba myosin II tail (Ch-Ac) was described in our previous paper (Liu et al., 2000Go). The Dictyostelium myosin II head and chicken gizzard smooth muscle myosin II tail chimera (Ch-Sm) was constructed as follows. The entire tail cDNA was amplified by PCR and fused with NcoI and HindIII sites at the 5' and 3' ends using pT7-7, which was kindly provided by Mitsuo Ikebe, University of Massachusetts, as the template. The primers were TGGCCATGGTGGAAACTCTTCTCAAAGGCTCGTCCACTGCTACAGGTCACCC and ATTGAATTCGGATCCTTAAGCTTCACTGGCTTTTCCATTGA. The product was subcloned into Litmus 28 and the sequences near both ends were verified: from the 5' end to the first NsiI site and from the 3' end to the EcoO66 site. The sequence between the first NsiI and EcoO66 sites was replaced by the native fragment of pT7-7 to eliminate potential PCR-derived point mutations within this region. The resultant NcoI-HindIII fragment was used to replace the corresponding Dictyostelium tail sequence. The resultant construct consisted of Dictyostelium myosin head from the N-terminus to residue Arg877 fused to chicken gizzard smooth muscle myosin tail beginning with Pro849 of smooth muscle myosin (which became Pro878 of Ch-Sm) with the C-terminal Glu replaced by Ala for fusion of the restriction site. The GFP-Dd-Wt construct was as described (Yumura and Uyeda, 1997aGo; Yumura and Uyeda, 1997bGo) and GFP-Ch-Sm and GFP-Ch-Ac were prepared with minor modifications of the same procedure.

For construction of Acanthamoeba myosin II tail cDNA, PCR was used to synthesize the actin 15 promoter using pMyD as the template, creating an EcoRV site directly downstream the initial codon. The PCR product was subcloned into Litmus 29 by XbaI and EcoRV resulting in Lit29A15, confirmed by sequencing. A fragment containing an NcoI site at the 5' end and a SacI site at the 3' end was synthesized by PCR using pMyD as the template, creating a stop codon and an MluI site directly downstream the stop codon at the 3' end. The PCR product was subcloned into Lit29A15 by SacI and NcoI, resulting in Lit29A15Mlu. The tail of Acanthamoeba myosin II liberated from pTIKLC-AC (Liu et al., 2000Go) by digestion with EcoRV and MluI was subcloned into Lit29A15Mlu. The resultant tail cDNA was subcloned into the expression vector pTIKL C-Ac by restriction with MluI and XbaI.

Cell culture
Wild-type AX3 cells and HS1 cells that lacked the endogenous copy of myosin heavy chain (Ruppel et al., 1994Go) were cultured on plastic plates and all suspension cultures were grown in HL5 medium supplemented with 60 µg/ml of penicillin and streptomycin at 21°C (Sussman, 1987Go). Dd-Wt, Ch-Ac and Ch-Sm heavy chain cDNAs and Acanthamoeba myosin II tail cDNAs were electroporated into HS1 cells using a Bio-Rad Laboratories (Hercules, CA) gene pulser (Egelhoff et al., 1991bGo). Individual clones were selected and maintained in the presence of 12 µg/ml geneticin (G418) sulfate (Life Technologies, Great Island, NY) in HL5 medium containing 60 µg/ml each of penicillin and streptomycin.

Electrophoresis and western blots
Cells were harvested, washed with 10 mM Tris buffer, pH 7.5, lysed with Triton X-100 and centrifuged for 1 hour at 240,000 g in a Beckman TL100 centrifuge. Equivalent amounts of the pellets were loaded onto 7.5% SDS-PAGE (Laemmli, 1970Go). Gels were either stained with Coomassie blue or transferred electrophoretically to nitrocellulose paper. Blots were reacted with three different antibodies: anti-Dictyostelium myosin II, anti-Acanthamoeba myosin II and anti-human nonmuscle myosin II (Fujiwara and Pollard, 1976Go), which recognizes the Ch-Sm heavy chain.

To analyze the time course of expression of cAMP receptor cAR1, cells were washed in developmental buffer, 5 mM Na2HPO4, 5 mM KH2PO4, 2 mM MgSO4, 200 µM CaCl2, pH 6.2, and resuspended in 5 ml of the same buffer at 2x107 cells/ml. The cells were pulsed with 75 nM cAMP (final concentration) at 6 minute intervals and 100 µl samples were taken at different times, lysed with 5x concentrated SDS sample buffer and subjected to SDS-PAGE. The gel was probed with anti-cAR1 antibody (Kim et al., 1998Go) followed with horseradish peroxidase-conjugated goat anti-rabbit antibody (Bio-Rad, Richmond, CA).

Growth rate assay
Cells were grown in 50 ml conical flasks on a rotary shaker at 175 rpm at 21°C (Sussman, 1987Go). Cell numbers were counted daily using a hemocytometer. After 3 days in suspension culture, cells were fixed and stained with 1 µg/ml of 4,6-diamidino-2-phenylindole.

Capping of ConA receptors and immunomicroscopy
To determine the time course of capping, cells were washed twice and resuspended in starvation buffer, 20 mM morpholinoethanesulfonic acid, pH 6.8 (Pasternak et al., 1989aGo; Egelhoff et al., 1991bGo). A small drop of cells was applied to glass coverslips and, after the cells attached, 50 µl of 30 µg/ml tetramethyl rhodamine isothiocyanate-conjugated ConA (Molecular Probes, Eugene, OR) were added and the cells were incubated for various times. Excess ConA was removed by rinsing the coverslips briefly in starvation buffer. The cells were fixed and permeablized in -20°C acetone plus 1% formalin for 15 minutes (LeBlanc-Straceski et al., 1994Go), the coverslips were washed and then either mounted in glycerol directly to visualize the ConA caps or reacted with anti-myosin antibodies for co-localization of myosin.

For localization of myosin during the cell cycle by indirect immunofluorescence, cold treatment was employed, as described previously (Shu et al., 1999Go), to partially synchronize the cell cycle. Cells were incubated with the same primary antibodies used in the Western blot analyses; anti-Dictyostelium myosin antibody was diluted 1:1000 and anti-Ch-Ac and anti-Ch-Sm antibodies were diluted 1:100. Fluorosceine isothiocyanate-conjugated goat anti-rabbit IgG (Molecular Probes) was used at 1:150 dilution. Micrographs were taken on a Zeiss LSM-510 laser scanning fluorescence microscope equipped with a Plan apo 63x oil objective.

Cell streaming
Streaming assays were performed as described (Peterson et al., 1995Go), with minor modification. Briefly, cells were harvested at mid-log phase, resuspended at 5x106/ml and 1.5x107 cells were plated on 60 mm petri dishes and allowed to adhere for 30 minutes. The medium was aspirated off, the cells washed with starvation buffer, 2 ml of starvation buffer were carefully applied and the cells placed in the dark for 6-8 hours.

Phagocytosis
The rate of uptake of fluorosceine isothiocyanate-conjugated, 1 µm polystyrene-latex beads (Polysciences, Warrington, PA) was quantified as described (Vogel et al., 1980Go; Jung et al., 1996Go) with slight modifications. Cells were collected, washed in Sorenson's buffer (16 mM KH2PO4, 2 mM Na2HPO4, pH 6.1) and diluted to 4x106 cells/ml in 20 ml of the same buffer. Cells were shaken at 100 rpm for 30 minutes at room temperature, washed beads were added at a ratio of 300 beads per cell and 1 ml of cells was removed at different times and added to 2 ml of ice-cold Sorenson's buffer to stop phagocytosis. Uningested beads were removed by centrifugation through 10 ml of a 20% polyethyleneglycol cushion and the cell pellets were suspended in 3 ml of 50 mM Na2HPO4, pH 9.2, and lysed by addition of Triton X-100 to 0.4%. Fluorescence intensity was measured at excitation and emission wavelengths of 485 and 520 nm, respectively. Cell numbers were determined and the final fluorescence intensity was corrected for total protein concentration as determined by the Bio-Rad method.

Plaque expansion assay
This assay was performed as described (Nellen et al., 1987Go) with minor modifications. A mixture of Dictyostelium cells and heat-killed Klebsiella aerogenes was seeded onto Millipore black filters on pads saturated with starvation buffer in petri dishes and placed in the dark in a humid chamber 21°C. Plaque sizes were measured after 5 days.

Development assay
Cell development was assessed by spotting small aliquots of amoebae onto a lawn of Klebsiella aerogenes on SM/5 agar plates (Kubalek et al., 1992Go). Phenotypes were observed under a dissection microscope over a 48 hour period.

Cell motility and chemotaxis assays
Cells were grown in HL5 media to densities of ~5x106 cells/ml, harvested by centrifugation, and allowed to differentiate by resuspending them at 2x107 cells/ml in 5 mM Na2HPO4, 5 mM KH2PO4, pH 6.2, 2 mM MgSO4, 200 µM CaCl2 and shaking at 100 rpm for 5-8 hours with repeated pulses of 75 nM cAMP, final concentration of each pulse (Devreotes et al., 1987Go). Cells were harvested by centrifugation, resuspended in 5 mM Na2HPO4, 5 mM NaH2PO4, pH 6.2, plated in a small spot on a chambered coverslip (Lab-Tek, Nalge Nunc, Naperville, IL), allowed to adhere, and covered with the same buffer to avoid drying. The cells were visualized using an inverted Zeiss microscope (Axiovert 100, Thornwood, NY) equipped with 10 and 20x Plan A Varel objectives lenses. Cell movement was recorded using a Photometrics, Coolsnap PXL CCD camera controlled by the IPLab-Spectrum software using a G4 Macintosh computer. For the chemotaxis assays, the chemoattractant gradients were generated using an Eppendorf microinjector connected to micropipettes containing 10 µM cAMP, as previously described (Parent et al., 1998Go). Quantification analysis was performed using the Digital Image Analysis System (DIAS7) software (Solltech, Oakdale, IA).

Determination of cellular ATP concentration
ATP concentration was determined using the luciferase-luciferin assay (Sigma, St Louis, MO) according to the protocol provided by the manufacturer with slight modification. In brief, cells were lysed in passive lysis buffer (Promega, Madison, WI) by two rounds of freezing in dry ice and thawing. The lysate was centrifuged at 16,000 g for 10 min at 4°C. The supernatant was adjusted to a concentration equivalent to the lysate of 5x106 cells/ml and 20 µl of the adjusted supernatant were gently mixed with 100 µl of luciferase-luciferin. The fluorescence was immediately measured using a TD-2020 Luminometer (Turner Designs, Sunnyvale, CA).


    Results
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Expression of chimeric myosins and cell growth in suspension culture
Dd-Wt was expressed in transfected HS1 null cells (Fig. 1C,D) at about 80% the level of expression of DdMII in the parental AX3 cells from which the HS1 cell line was derived (Ruppel et al., 1994Go). On a molar basis, Ch-Ac was expressed at 80% and Ch-Sm at 41% the level of Dd-Wt expression (Fig. 1C). As revealed by immunofluorescence, all cells expressing Ch-Ac and Ch-Sm contained about the same amount of myosin and extended culture in suspension did not increase the level of either chimera (data not shown). Ch-Ac and Ch-Sm rescued growth (cytokinesis) of HS1 cells in suspension culture (Fig. 2A) to about the same extent as Dd-Wt (after a short lag phase for Ch-Sm), which was approximately 65% the rate of growth of parental AX3 cells.



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Fig. 2. Ch-Ac and Ch-Sm support growth and cytokinesis in suspension culture of transfected HS1 cells. (A) Growth rates of cells determined by cell number. (B) Representative cells stained with 4,6-diamidino-2-phenylindole after 3 days in suspension culture showing that transfected cells have many fewer nuclei than HS1 cells. (C) Quantification of number of nuclei per cell; the data are the mean of three experiments in each of which at least 100 cells were scored for each cell type.

 

Because HS1 cells cannot divide in suspension culture but mitosis is unaffected, surviving HS1 cells were very much larger than AX3 cells (Fig. 2B) and contained many more nuclei (Fig. 2B,C). HS1 cells expressing Dd-Wt were essentially the same size as AX3 cells with only slightly fewer mononucleate and binucleate cells and a few more cells with 3-10 or 11-30 nuclei (Fig. 2B,C). HS1 cells expressing Ch-Ac were similar in size to HS1 cells expressing Dd-Wt but with some increase in cells with more than 3 nuclei, whereas Ch-Sm cells were significantly larger and contained substantially more polynucleate cells. However, Ch-Sm cells were definitely smaller and had many fewer nuclei than HS1 cells (Fig. 2B,C). Thus, expression of either chimeric myosin rescued cytokinesis; Ch-Ac was almost as effective as Dd-Wt but Ch-Sm was less effective.

Localization of chimeric myosins in interphase and dividing cells
At interphase, Dd-Wt and Ch-AC expressed in HS1 cells had similar, predominantly cortical, localizations, like DdMII in AX3 cells, whereas Ch-Sm was predominantly concentrated in a single cortical patch (Fig. 3A) in both mono- and multi-nucleate cells; any Ch-Sm lying outside the patch was at too low a concentration to be detected by immunofluorescence. The position of the patch did not correlate with cell polarity (not shown). Despite their different localizations in interphase cells, however, both chimeras were concentrated at the cleavage furrow during anaphase and telophase (Fig. 3B,C) consistent with the ability of both chimeras to support growth and cytokinesis in suspension culture.



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Fig. 3. Localization of Ch-Sm differs from localization of Dd-Wt and Ch-Ac in interphase and dividing cells. (A) Interphase AX3 cells and HS1 cells expressing Dd-Wt, Ch-Ac and Ch-Sm. (B) Dividing cells. (C) HS1 cells expressing Ch-Sm at interphase, early telophase and late telophase. (D) HS1 cells expressing the tail of Acanthamoeba myosin II at interphase, anaphase and telophase. Myosins were localized by indirect immunofluorescence.

 

Zang and Spudich had shown that the actin-binding motor (head and neck) domain of DdMII is not required for localization of the tail domain to the cleavage furrow, implying that any required sequence for cleavage-furrow localization resides in the tail domain (Zang and Spudich, 1998Go). The results of Shu et al. imply further that if there is a required sequence it is probably within the relatively small region of the tail required for polymerization (Shu et al., 1999Go). The fact that the chimeric myosins with Acanthamoeba and smooth muscle myosin tails also localized to the cleavage furrow suggested the possibility that no specific sequence in either the motor (head and neck) or tail is required for localization.

To explore this possibility further, we expressed just the tail of Acanthamoeba myosin II in HS1 cells. At interphase, the expressed tail localized mostly in a cortical ring, like Dd-Wt and Ch-Ac (Fig. 3D), but with occasional patches. At anaphase and telophase, a substantial fraction of the expressed Acanthamoeba myosin tail (as judged by immunofluorescence) moved to the cleavage furrow leaving the cortical patches behind (Fig. 3D).

To follow the dynamics of myosin relocation in living cells, we expressed N-terminal GFP-labeled Dd-Wt in AX3 and HS1 cells and GFP-Ch-Ac and GFP-Ch-Sm in HS1 cells. The level of expression of GFP-Ch-Sm was too low to detect but time lapse images of the relocation of GFP-Dd-Wt and GFP-Ch-Ac from the cortex to the equatorial region and the constricting cleavage furrow, as illustrated in Fig. 4A for GFP-Ch-Ac, were as previously published for GFP-Dd-Wt (Yumura and Uyeda, 1997aGo; Zang and Spudich, 1998Go; Robinson et al., 2002Go). Because of the inability to express sufficient amounts of GFP-Ch-Sm, we localized Ch-Sm in dividing cells by indirect immunofluorescence and selected different cells whose images (Fig. 4B) most closely resembled the images of the GFP-Ch-Ac sequence. From the reconstructed sequence, it would appear that Ch-Sm did not redistribute to the cleavage furrow like Ch-Ac but that the cleavage furrow formed at the site of the cortical patch, which remained at one side of the cleavage furrow (consistent with the image of Ch-Sm in Fig. 3B) until the furrow became highly constricted.



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Fig. 4. Dynamics of localization of Ch-Ac and Ch-Sm to the cleavage furrow during cytokinesis. (A) Time course of localization of GFP-Ch-Ac to the cleavage furrow of a typical, live cell undergoing cytokinesis. Time is in seconds. (B) Gallery of images showing the localization by indirect immunofluorescence of Ch-Sm in cells at different stages of cytokinesis. The cells in B were aligned by matching them to similar images in the Ch-Ac sequence (A). Whereas Ch-Ac relocated symmetrically from the cortex to the cleavage furrow, the cleavage furrow in HS1 cells expressing Ch-Sm seemed to form at the site of the cortical patch of Ch-Sm, which remained on one side of the cleavage furrow until the furrow was highly constricted.

 

Chimeric myosins rescue capping of liganded ConA receptors
Within 5 minutes after ConA binds to its surface receptors in AX3 cells, the liganded receptors are transported to a surface cap with transient co-localization of myosin II to an underlying `cap' beneath the plasma membrane [Fig. 5 (Egelhoff et al., 1993Go)]; the myosin redisperses (Carboni and Condeelis, 1985Go) within approximately 10 minutes with no apparent change in the ConA cap [Fig. 5 (Egelhoff et al., 1993Go)]. ConA binds to the surface of HS1 cells but the receptors do not cap [(Egelhoff et al., 1993Go) and data not shown]. Capping of ConA receptors in HS1 cells expressing Dd-Wt was indistinguishable from capping in AX3 cells; ConA and Dd-Wt co-capped within 5 minutes (Fig. 5A) and the myosin `cap' was substantially dispersed by 10 minutes (Fig. 5B,C) and almost totally dispersed by 20-60 minutes (Fig. 5B,C). In HS1 cells expressing Ch-Ac and Ch-Sm, ConA caps and myosin co-caps also formed within 5 minutes (Fig. 5A). However, myosin dispersal was very much slower for Ch-Ac, only 40% dispersion by 60 minutes and 80% dispersion by 90 minutes (Fig. 5B,C). The Ch-Sm co-cap did not disperse even after 90 minutes (Fig. 5B,C); in fact, ConA and Ch-Sm remained co-localized even in ConA-capped cells undergoing cytokinesis (data not shown).



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Fig. 5. Ch-Ac and Ch-Sm rescue ConA-capping but Ch-Sm does not redisperse after capping. Distribution of ConA and myosins 5 minutes (A) and 20 minutes (B) after initiating capping by addition of ConA. (C) Percentage of cells with myosin caps as a function of time after initiating capping by addition of ConA. The ConA caps did not disperse in any of the cell lines. ConA, red; myosins, green.

 

Chimeric myosins partially rescue plaque expansion and Ch-Ac rescues mildly impaired phagocytosis
When Dictyostelium cells are plated on a lawn of bacteria they phagocytose the bacteria and, as the amoebae divide and move, form an expanding, bacteria-free plaque. The greatly inhibited plaque expansion activity of HS1 cells compared to AX3 cells (Fig. 6A) was essentially totally rescued by expression of Dd-Wt (Fig. 6A), rescued somewhat less well by Ch-Ac (Fig. 6A) and only partially rescued by Ch-Sm (Fig. 6A). To test whether the rates of plaque expansion might be related to rates of phagocytosis, which had been reported to occur in myosin II-null cells (De Lozanne and Spudich, 1987Go) but had not been quantified at the time these experiments were carried out, we measured the rates of phagocytosis of fluorescently labeled latex beads (Fig. 6B). When calculated on the basis of total cell protein, phagocytosis by HS1 cells was about 25% inhibited compared to AX3 cells (Fig. 6B). This inhibition was reversed by expression of Dd-Wt and Ch-Ac but apparently not by expression of Ch-Sm (Fig. 6B). From these data it seems unlikely that the differences in rates of plaque expansion were due solely, if at all, to impaired phagocytosis.



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Fig. 6. Plaque expansion and phagocytosis assays. (A) The deficiency of HS1 cells in the plaque expansion assay was largely reversed by expression of Dd-Wt, less well by expression of Ch-Ac and only slightly by expression of Ch-Sm. Seven to ten drops of Dictyostelium amoebae in a suspension of heat-killed bacteria were spotted onto black filters and micrographs were taken 5 days later when the expanding amoeba plaques had ingested the bacteria and exposed the black filter. (B) Phagocytosis of latex beads was only 25% inhibited in HS1 cells and was completely rescued by expression of Dd-Wt and Ch-Ac. HS1 cells expressing Ch-Sm were no different than HS1 cells. Each point is the mean of five independent experiments for AX3 and three independent experiments for the other cell lines. Standard deviations were about 10% for the later time points in experiments with AX3, Dd-Wt and Ch-Ac, and 20-30% for HS1 and Ch-Sm. Uptake was based on protein concentration rather than cell number to compensate for any differences in cell size. Cells expressing Ch-Sm were larger than the other cell lines, which were all about the same size.

 

Chimeric myosins partially rescue motility and chemotaxis
When Dictyostelium amoebae are plated on a surface in the absence of nutrients, they enter a developmental program that allows them to come together by chemotaxing towards secreted cAMP and, by streaming in a head-to-tail fashion, the cells form aggregation centers that eventually differentiate into mounds. Cell motility can be assessed by measuring this streaming response. Myosin II is not required for cell streaming (Eliott et al., 1993Go) but HS1 cells form broader more slowly moving streams culminating in flatter mounds than AX3 cells (Fig. 7A). Expression of Dd-Wt in HS1 cells rescued wild-type behavior (Fig. 7A). However, HS1 cells expressing either Ch-Ac or Ch-Sm did not form streams but moved as individual cells for short distances thereby forming smaller aggregates than AX3 or HS1 cells (Fig. 7A). In addition, HS1 cells expressing Ch-Ac or Ch-Sm appeared to be rounder than either AX3 or HS1 cells.



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Fig. 7. Assays of cell streaming and chemotaxis. (A) In the cell streaming assay, micrographs were taken 7-8 hours after plating the amoebae on non-nutrient medium. (B) In the chemotaxis assay, cells were pulsed with 75 nM cAMP (final concentration after each pulse) for the time required for maximum expression of cAMP receptors (see text), washed and allowed to adhere to a coverslip, and photographed at 15-second intervals. The micropipette contained 10 µM cAMP.

 

Another way to assess motility of Dictyostelium amoebae is to study chemotaxis towards a point source of cAMP. For these experiments, in order for the cells to express the signaling machinery required to sense the chemoattractant cAMP, cells are suspended in non-nutrient buffer and pulsed with cAMP every 6 minutes (see Materials and Methods). The extent of development is monitored by measuring the appearance of the cAMP receptor, cAR1, by Western analysis. When AX3, HS1 and HS1 cells expressing Dd-Wt were pulsed with cAMP, cAR1 increased to a maximum at about 5-6 hours (data not shown). In contrast, the expression of cAR1 in HS1 cells transformed with either Ch-Ac or Ch-Sm reached maximum levels at about 8 hours (data not shown). Perhaps importantly, the maximum concentration of cAR1 in HS1 cells expressing Ch-Sm was less than in HS1 cells.

With these data in mind, we measured chemotaxis after 6 hours of cAMP pulsing for AX3, HS1 and HS1 cells expressing Dd-Wt and after 8 hours for HS1 cells expressing either Ch-Ac or Ch-Sm. The chemotaxis competent cells were plated on chambered cover slips and the cAMP gradient was established using a micropipette. Fig. 7B shows frames taken just before and 6 minutes after the cAMP gradient was established for the four cell lines. The velocity of chemotaxis, directional change and percent roundness were quantified by computer analysis of movies taken over a period of 10 minutes with frames at 15 second intervals. As shown in Table 1, both the reduced rate of chemotaxis towards a point source of cAMP and the greater directional change by HS1 cells compared to AX3 cells were completely rescued by expression of Dd-Wt. However, Dd-Wt only partially rescued the elongated shape of AX3 cells. On the other hand, expression of Ch-Ac and Ch-Sm in HS1 cells only partially rescued chemotactic motility and, surprisingly, both directional change and roundness of the cells expressing Ch-Ac and Ch-Sm were no different than for HS1 cells. In all cases, the myosin was concentrated in the posterior region of the migrating cells (Fig. 8) as expected (Fukui et al., 1989Go).


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Table 1. DIAS® analysis of chemotaxing Dictyostelium

 


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Fig. 8. Ch-Ac and Ch-Sm concentrate in the posterior region of migrating cells. Cells were pulsed with 75 nM cAMP (final concentration) as in Fig. 9. At the times of maximum cAR1 concentration (see text), cells were washed and placed on a coverslip and allowed to migrate towards a pipette containing 10 µM cAMP, as in Fig. 7. Micrographs of cells expressing GFP-myosins were taken while the cells were under observation so the direction of movement was known. The localization of Ch-Sm was determined by indirect immunofluorescence of fixed cells and the direction of movement inferred from the orientation of the pseudopods. The cells shown are typical. All were moving from right to left. GFP-Dd-Wt-AX3, AX3 cell expressing GFP-Dd-Wt; GFP-Dd-Wt, HS1 cell expressing GFP-Dd-Wt; GFP-Ch-Ac, HS1 cell expressing GFP-Ch-Ac; Ch-Sm, HS1 cell expressing Ch-Sm.

 



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Fig. 9. Ch-Ac and Ch-Sm do not support full development to fruiting bodies. AX3 cells developed to fruiting bodies; HS1 never progressed beyond the mound stage; Dd-Wt rescued full development in HS1 cells; HS1 cells expressing Ch-Ac did not develop beyond the mound stage; Ch-Sm-transfected HS1 cells developed a few aberrant fingers but never developed to fruiting bodies. All micrographs were taken at 48 hours after plating the amoebae.

 
Chimeric myosins do not rescue development
When starved, AX3 cells move in streams aggregating to form mounds that differentiate into multicellular slugs that migrate for approximately 6 hours before differentiating to form fingers that finally develop into well-differentiated stalks and spore-containing fruiting bodies by 24 hours. Cells not expressing myosin are unable to develop past the mound stage (Knecht and Loomis, 1987Go; De Lozanne and Spudich, 1987Go; Manstein et al., 1989Go). Expression of Dd-Wt in HS1 cells rescued essentially normal development to fruiting bodies (Fig. 9). However, HS1 cells expressing Ch-Ac formed abnormal mounds that did not progress further while HS1 cells expressing Ch-Sm developed to the finger stage but never beyond (Fig. 9).

We expressed Ch-Ac in AX3 cells to determine whether chimeric myosins would inhibit normal development. Development of AX3 was unaffected when Ch-Ac was expressed at 34% the level of DdMII, about half of the level of expression of Ch-Ac in HS1 cells but similar to the expression level of Ch-Sm in HS1 cells (41% of Dd-Wt). Because the myosins in transfected HS1 cells were extrachromosomally expressed under control of the actin15 promoter, we thought their expression might be regulated differently during development than the expression of endogenous myosin. We found, however, that the amounts of Dd-Wt and Ch-Ac in HS1 cells and DdMII in AX3 cells were unchanged after 6 hours in starvation medium under the conditions used to follow development. Because the chimeras have greatly enhanced ATPase activity in vitro, we determined the ATP levels (by luciferinase-luciferin assay) during development and found them to be 35% lower at 6 hours in both AX3 cells and HS1 cells expressing Ch-Ac.


    Discussion
 Top
 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We have shown that expression of chimeras consisting of the motor domain of Dictyostelium myosin II fused to the tail domain of either Acanthamoeba or chicken gizzard smooth muscle myosin II restores cytokinesis and growth in suspension culture of myosin heavy chain-null cells (HS1) almost as well as expression of wild-type myosin heavy chain (but Ch-Sm not quite as well as Ch-Ac), that both chimeras and even Acanthamoeba myosin II tail alone localize to the cleavage furrow like wild-type myosin, that both chimeras restore the ability to cap ConA receptors but dissociate from the ConA cap more slowly than Dd-Wt (Ch-Sm apparently not at all), that the reduced rate of chemotaxis of HS1 cells is partially restored by both chimeras, and that the minor impairment of phagocytosis of latex beads by HS1 cells is completely restored by Ch-Ac but not at all by Ch-Sm.

When cells are pulsed with cAMP, cAMP receptors increase more slowly, and to a lesser extent, and decrease more slowly in HS1 cells expressing the chimeric myosins than in HS1 cells expressing Dd-Wt or even in non-transfected HS1 cells. Interestingly, in the cell-streaming assay HS1 cells expressing either Ch-Ac or Ch-Sm do not form obvious streams but expression of Dd-Wt rescues HS1 streaming to the level of AX3 cells. In the plaque expansion assay, Dd-Wt and Ch-Ac almost completely rescue HS1 cells but Ch-Sm rescues only slightly. Importantly, although development is completely rescued by Dd-Wt, neither chimera rescues the inability of HS1 cells to develop into stalks and fruiting bodies.

We are aware of only two experiments similar to those described in this paper. In one (Shu et al., 1999Go), a chimera of the Dictyostelium motor domain and chicken skeletal muscle myosin tail expressed in myosin II null cells localized to the cleavage furrow and supported cytokinesis. The ability of this chimera to support other functions of DdMII was not tested. In the other (LeBlanc-Straceski et al., 1994Go) a chimera consisting of the Dictyostelium motor domain and ß-cardiac myosin subfragment 2 with the light meromyosin region (the region essential for filament formation) was expressed in wild-type and myosin II heavy chain-null cells. The ß-cardiac chimera, which was expressed in null cells to only about 8% the level of DdMII in wild-type cells, occurred in either a uniformly dispersed punctate pattern or in a cortical patch not unlike that observed by us for Ch-Sm. The ß-cardiac chimera rescued capping of ConA receptors but, like Ch-Sm in our experiments, did not redisperse from the co-cap after 45 minutes. Unlike Ch-Ac and Ch-Sm in our experiments, the ß-cardiac chimera did not support growth in suspension culture, possibly because its expression level was too low (Egelhoff et al., 1991aGo). The ability of the ß-cardiac myosin chimera to localize to the cleavage furrow of cells undergoing cytokinesis on a substrate was not determined. Like Ch-Ac and Ch-Sm, the ß-cardiac chimera did not support development past the mound stage. The biochemical properties of the ß-cardiac chimera were not characterized by in vitro studies.

Myosin II mutants with deletions of tail residues 824-941, 943-1194, 943-1464 or 1156-1464 (Fig. 10) all move to the cleavage furrow (Kubalek et al., 1992Go; Shu et al., 1999Go) and myosin II truncated at residue 1818 supports cytokinesis in suspension culture whereas myosin truncated at residue 1783 does not (Lee et al., 1994Go). Taken together, these data suggest that the only region required for localization to the cleavage furrow lies between tail residues 1465 and 1818. Interestingly, the minimum expressed peptide shown to polymerize in vitro similarly to full-length DdMII, residues 1533-1819 (O'Halloran et al., 1990Go) lies within this region.



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Fig. 10. Alignment of tail sequences of Acanthamoeba, Dictyostelium and chicken smooth muscle myosins reveals limited similarities. Sequences beginning with the invariant Pro that defines the head/tail junction (Korn, 2000Go) were aligned by the default mode of the PC version of Clustal X (Thompson et al., 1997Go). Amino acids that occur at the same positions in all three myosins are in red, those that occur only in Acanthamoeba and Dictyostelium myosins are in blue, and those that occur in smooth muscle and Dictyostelium myosins are in green. The region of highest similarity in the three myosins and the corresponding region of chicken skeletal muscle myosin are enlarged at the bottom of the figure. Amino acids in skeletal muscle myosin that are the same as those in the corresponding positions of the three other myosins are in red, those that are the same as in Acanthamoeba and Dictyostelium myosins are in blue, those that are the same as in smooth muscle and Dictyostelium myosins are in green and those that are the same only in skeletal and Dictyostelium myosins are in purple. The GenBank accession numbers are: Acanthamoeba myosin, P05659; Dictyostelium myosin, A26655; smooth muscle myosin, P01587; skeletal muscle myosin, P13538.

 

The tail sequences of Dictyostelium, Acanthamoeba and chicken smooth muscle myosins are quite different (Fig. 10) which limits the possibilities for any specific sequence that might be required for localization of myosin II to the cleavage furrow. The region of highest sequence similarity between Acanthamoeba, smooth muscle and Dictyostelium myosin II tails (Fig. 10) corresponds to Dictyostelium residues 1695-1741 (Acanthamoeba residues 1304-1350, smooth muscle residues 1534-1580), which lie within the maximal regions required for cleavage furrow localization and polymerization of expressed peptides. Thus, if there is a specific tail sequence required for localization of DdMII to the cleavage furrow it probably lies within these 47 amino acids which, for all three myosins, have a 100% probability of being in a coiled-coil conformation, according to Paircoil (Berger et al., 1995Go), beginning with residue b in the first heptad and ending with residue f in the last heptad (data not shown). The only significant sequence within that region that is shared by all three myosins is Dictyostelium residues 1714-1720 (QLEEEED) which corresponds to Acanthamoeba residues 1323-1329 (QLEEEQD) and smooth muscle residues 1553-1559 (QLEELED). Possibly this sequence is required for localization to the cleavage furrow. However, a myosin II chimera with the tail of chicken skeletal muscle myosin, in which the corresponding sequence (ALEEAEA, see Fig. 10, bottom) is less similar to the Dictyostelium sequence also localizes to the cleavage furrow (Shu et al., 1999Go). Therefore, it seems quite possible that no specific sequence is required for localization of myosin II to the cleavage furrow of Dictyostelium and that myosin filaments are transported to the cleavage furrow by an unknown mechanism that requires neither myosin II motor activity, nor specific interaction of myosin II with actin filaments nor specific, high-affinity interactions with any other protein.

We had hoped to follow the dynamics of GFP-labeled Ch-Sm in live cells to determine whether Ch-Sm moved from the dense cortical patch to the cleavage furrow or if the cleavage furrow formed at the position of the patch. Although too little GFP-Ch-Sm was expressed to do this experiment, reconstruction of the dynamics by comparing indirect immunofluorescence images of HS1 cells expressing Ch-Sm to dynamic images of HS1 cells expressing GFP-Ch-Ac indicates that the cleavage furrow formed at the location of an `immobile' Ch-Sm cortical patch. This conclusion is supported by the observation that in dividing, ConA-capped cells the ConA cap remained co-localized with the Ch-Sm cortical patch throughout cytokinesis. As only about 10% of myosin II concentrates at the cleavage furrow of wild-type cells (Robinson et al., 2002Go), the ability of Ch-Sm to support cytokinesis of HS1 cells in suspension culture might reflect just the activity of a small amount of cortical Ch-Sm lying outside the predominant cortical patch. In any case, if the filaments formed in vivo by Ch-Sm resemble the side-polar filaments formed in vitro, these experiments show that bipolar filaments are not required to support cytokinesis.

The slow dispersion of Ch-Ac compared to Dd-Wt following capping of ConA receptors may be due to the fact that, in contrast to Dd-Wt, depolymerization of Ch-Ac is not regulated. In this regard, Ch-Ac behaves similarly to the slowly dispersing point mutants (Egelhoff et al., 1993Go) and truncated mutant (Egelhoff et al., 1991aGo) of DdMII that lack the regulatory phosphorylation sites. The total inability of Ch-Sm to redisperse may result from the absence of regulated depolymerization accentuated by the physical properties of the large, side-polar filaments that probably also are responsible for the localization of Ch-Sm in interphase cells predominantly to a single, dense patch. The ability of Ch-Sm, despite its predominant localization to a single patch, to cap ConA receptors may be due to a small amount of dispersed myosin, too little to be detected by immunofluorescence but sufficient to pull the ConA receptors into the pre-existing patch that contains most of the myosin. In this case, Ch-Sm would not be `co-capping' with the ConA and one would, therefore, not expect the Ch-Sm `cap' (actually the pre-existing cortical patch) to disperse. The inability to express GFP-labeled Ch-Sm in detectable amounts made it impossible to evaluate this hypothesis in live cells.

The inability of HS1 cells expressing either of the chimeric myosins to form streams, in contrast to HS1 null cells, is unlikely to be due to an inherently greater rate of motility of HS1 cells because, in the chemotaxis assay (Table 1), HS1 cells move more slowly than HS1 cells expressing Ch-Ac or Ch-Sm. The inability to form streams may have resulted from either a lower concentration of cAMP receptors in the HS1 cells expressing chimeric myosins than in HS1-null cells (as observed when cells in suspension were pulsed with cAMP), a shallower gradient of cAMP (which, in the streaming assay is created by cells secreting cAMP) which was not measured, or a combination of these and perhaps other factors. On the other hand, the relative activities of the cells lines in the plaque expansion assay, HS1<Ch-Sm<Ch-Ac<Dd-Wt=AX3, are consistent with their relative motilities in the chemotaxis assay, HS1<Ch-Sm=Ch-Ac<Dd-Wt=AX3 (Table 1).

While Dd-Wt fully rescued the defect in chemotaxis of HS1 cells, chemotaxis was only partially rescued by expression of Ch-Ac or Ch-Sm. This partial rescue may have been due to the lower concentration of cAR1 receptors in HS1 cells expressing the chimeric myosins. The mechanism by which the chimeric myosins interfere with the expression of cAR1 receptors is an interesting future study. It may also be relevant to the partial rescue of chemotaxis by the chimeric myosins that, despite their very much higher actin-activated MgATPase activity, the in vitro motility activities of the chimeras are less than that of Dd-Wt (Liu et al., 2000Go).

Myosin II appears to make only a minor (possibly indirect) contribution to phagocytosis in as much as HS1 cells were about 75% as active as AX3 cells in our experiments and a recent paper by Maselli et al. reported no inhibition of phagocytosis in a different myosin-null cell line (Maselli et al., 2002Go). This is consistent with reports (Titus, 1999Go; Tuxworth et al., 2001Go) that myosin VII has a major role in phagocytosis; the initial rate of phagocytosis by myosin VII-null cells is only 10% the rate of control cells (Tuxworth et al., 2001Go). The apparent inability of Ch-Sm to rescue the minor impairment of phagocytosis in myosin II-null cells may be a result of most of the Ch-Sm being concentrated in the cortical patch leaving too little to contribute, whether directly or indirectly, to the phagocytic process.

Why are the chimeric myosins unable to support full development? It cannot be because they are extrachromosomally expressed because Dd-Wt expressed in the same plasmid does support full development of HS1 cells and the level of Ch-Ac, like the level of Dd-Wt and DdMII in AX3 cells, did not change during at least 6 hours of development in starvation medium. Also, it is unlikely to be due to the fact that neither the actin-activated MgATPase activity nor filament formation of the chimeras is regulated because similarly unregulated point mutants of myosin II support full development (O'Halloran and Spudich, 1990Go; Uyeda and Spudich, 1993Go; Ostrow et al., 1994Go; Chen et al., 1999Go). Although the ATPase activities in vitro of the chimeric myosins are much higher than the activity of DdMII, the levels of ATP were similar in AX3 and HS1 cells expressing Ch-Ac after 6 hours in development medium and expression of Ch-Ac in AX 3 cells did not affect their full development to fruiting bodies. Therefore, an insufficiency of ATP is probably not the reason the chimeric myosins cannot support development of HS1 cells.

Springer et al. showed that myosin II is required at two stages of development: first, during aggregation for cells to sort properly and, again, near the end of development, for completion of stalk formation and positioning of the fruiting body (Springer et al., 1994Go). HS1 cells expressing Ch-Ac and Ch-Sm appear to be blocked at the first myosin II-dependent stage. Apparently, the properties of the chimeric myosin filaments render them unable to function properly in the early stages of multicellular development.


    Acknowledgments
 
We thank Keigi Fujiwara, University of Rochester, and Kohji Ito, Chiba University, for generously providing antibodies to smooth muscle myosin heavy chain and Dictyostelium myosin II heavy chain, respectively.


    References
 Top
 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

Aguado-Velasco, C. and Bretscher, M. S. (1997). Dictyostelium myosin II null mutant can still cap ConA receptors. Proc. Natl. Acad. Sci. USA 94, 9684-9686.[Abstract/Free Full Text]

Berger, B., Wilson, D. B., Wolf, E., Tonchev, T., Milla, M. and Kim, P. S. (1995). Predicting coiled coils by use of pairwise residue correlations. Proc. Natl. Acad. Sci. USA 92, 8259-8263.[Abstract]

Berlot, C. H., Spudich, J. A. and Devreotes, P. N. (1985). Chemoattractant-elicited increases in myosin phosphorylation in Dictyostelium. Cell 43, 307-314.[CrossRef][Medline]

Berlot, C. H., Devreotes, P. N. and Spudich, J. A. (1987). Chemoattractant-elicited increases in Dictyostelium myosin phosphorylation are due to changes in myosin localization and increases in kinase activity. J. Biol. Chem. 262, 3918-3926.[Abstract/Free Full Text]

Carboni, J. M. and Condeelis, J. S. (1985). Ligand-induced changes in the location of actin, myosin, 95K (alpha actinin) and 120K protein in amebae of Dictyostelium discoideum. J. Cell Biol. 100, 1884-1893.[Abstract]

Chen, P., Chaudoir, B. M., Trybus, K. M. and Chisholm, R. L. (1999). Expression of chicken gizzard RLC complements the cytokinesis and developmental defects of Dictyostelium RLC null cells. J. Musc. Res. Cell Motil. 20, 177-186.[Medline]

Craig, R. and Megerman, J. (1997). Assembly of smooth muscle myosin into side-polar filaments. J. Cell Biol. 75, 990-996.[Abstract]

De Lozanne, A. and Spudich, J. A. (1987). Disruption of the Dictyostelium myosin heavy chain gene by homologous recombination. Science 236, 1086-1091.[Medline]

De Lozanne, A., Lewis, M., Spudich, J. A. and Leinwand, L. A. (1985). Cloning and characterization of a nonmuscle myosin heavy chain cDNA. Proc. Natl. Acad. Sci. USA 82, 6807-6810.[Abstract]

Devreotes, P. N., Fontana, D., Klein, P., Sherring, J. and Theibert, A. (1987). Transmembrane signaling in Dictyostelium. Methods Cell Biol. 28, 299-331.[Medline]

Egelhoff, T. T., Brown, S. S. and Spudich, J. A. (1991a). Spatial and temporal control of nonmuscle myosin localization: identification of a domain that necessary for myosin filament formation in vivo. J. Cell Biol. 112, 677-688.[Abstract]

Egelhoff, T. T., Lee, R. J. and Spudich, J. A. (1991b). Spatial and temporal control of nonmuscle myosin localization: identification of a domain that is necessary for myosin filament disassembly in vivo. J. Cell Biol. 112, 677-688.[Abstract]

Egelhoff, T. T., Lee, R. J. and Spudich, J. A. (1993). Dictyostelium myosin heavy chain phosphorylation sites regulate myosin filament assembly and localization in vivo. Cell. 75, 363-371.[Medline]

Eliott, S., Joss, G. H., Spudich, J. A. and Williams, K. L. (1993). Patterns in Dictyostelium discoideum: the role of myosin II in the transition from the unicellular to the multicellular phase. J. Cell Sci. 104, 457-466.[Abstract/Free Full Text]

Fujiwara, K. and Pollard, T. D. (1976). Fluorescent antibody localization of myosin in the cytoplasm, cleavage furrow and mitotic spindle of human cell. J. Cell Biol. 71, 848-875.[Abstract]

Fukui, Y., Lynch, T. J., Brzeska, H. and Korn, E. D. (1989). Myosin I is located at the leading edges of locomoting Dictyostelium amoebae. Nature 341, 328-331.[CrossRef][Medline]

Fukui, Y., de Lozanne, A. and Spudich, J. A. (1990). Structure and function of the cytoskeleton of a Dictyostelium myosin-defective mutant. J. Cell Biol. 110, 367-378.[Abstract]

Griffith, L. M., Downs, S. M. and Spudich, J. A. (1987). Myosin light chain kinase and myosin light chain phosphatase from Dictyostelium: effects of reversible phosphorylation on myosin structure and function. J. Cell Biol. 104, 1309-1323.[Abstract]

Jung, G., Wu, X. and Hammer, J. A., III (1996). Dictyostelium mutants lacking multiple classic myosin I isoforms reveal combinations of shared and distinct functions. J. Cell Biol. 133, 305-323.[Abstract]

Kim, J. Y., Borleis, J. A. and Devreotes, P. N. (1998). Switching of chemoattractant receptors programs development and morphogenesis in Dictyostelium: receptor subtypes activate common responses at different agonist concentrations. Dev. Biol. 197, 117-128.[CrossRef][Medline]

Knecht, D. A. and Loomis, W. F. (1987). Antisense RNA inactivation of myosin heavy chain gene expression in Dictyostelium discoideum. Science 236, 1081-1086.[Medline]

Korn, E. D. (2000). Coevolution of head, neck, and tail domains of myosin heavy chains. Proc. Natl. Acad. Sci. USA 97, 12559-12564.[Abstract/Free Full Text]

Kubalek, E. W., Uyeda, T. Q. P. and Spudich, J. A. (1992). A Dictyostelium myosin lacking a proximal 58-kDa portion of the tail is functional in vitro and in vivo. Mol. Biol. Cell 3, 1455-1462.[Abstract]

Kuczmarski, E. R. and Spudich, J. A. (1980). Regulation of myosin self assembly: phosphorylation of Dictyostelium heavy chains inhibits formation of thick filaments. Proc. Natl. Acad. Sci. USA 77, 7292-7296.[Abstract]

Kuczmarski, E. R., Tafuri, S. R. and Parysek, L. M. (1987). Effect of heavy chain phosphorylation on the polymerization and structure of Dictyostelium myosin filaments. J. Cell Biol. 105, 2989-2997.[Abstract]

Laemmli, U. K. (1970). Cleavage of structural proteins during assembly of the head of bacteriophage T4. Nature 227, 680-685.[Medline]

LeBlanc-Straceski, J. M., Fukui, Y., Sohn, R. L., Spudich, J. A. and Leinwand, L. A. (1994). Functional analysis of a cardiac myosin rod in Dictyostelium discoideum. Cell Motil. Cytoskel. 27, 313-326.[Medline]

Lee, R. J., Egelhoff, T. T. and Spudich, J. A. (1994). Molecular genetic truncation analysis of filament assembly and phosphorylation domains of Dictyostelium myosin heavy chain. J. Cell Sci. 107, 2875-2886.[Abstract/Free Full Text]

Liang, W., Warrick, H. M. and Spudich, J. A. (1999). A structural model for phosphorylation control of Dictyostelium myosin II thick filament assembly. J. Cell Biol. 147, 1039-1048.[Abstract/Free Full Text]

Liu, X., Morimoto, S., Hikkoshi-Iwane, A., Yanagida, T. and Uyeda, T. Q. P. (1998). Filament structure as an essential factor for regulation of Dictyostelium myosin by regulatory light chain phosphorylation. Proc. Natl. Acad. Sci. USA 24, 14124-14129.

Liu, X., Shu, S., Yamashita, R. A., Xu, Y. and Korn, E. D. (2000). Chimeras of Dictyostelium myosin II head and neck domains with Acanthamoeba or chicken smooth muscle myosin II tail domain have greatly increased and unregulated actin-dependent MgATPase activity. Proc. Natl. Acad. Sci. USA 97, 12553-12558.[Abstract/Free Full Text]

Luck-Vielmetter, D., Schleicher, M., Grabatin, B., Wippler, J. and Gerisch, G. (1990). Replacement of threonine residues by serine and alanine in a phosphorylatable heavy chain fragment of Dictyostelium myosin II. FEBS Lett. 269, 239-243.[CrossRef][Medline]

Malchow, D., Böhme, R. and Rahmsdorf, H. J. (1981). Regulation of phosphorylation of myosin heavy chain during chemotactic response of Dictyostelium cells. Eur. J. Biochem. 117, 213-218.[Abstract]

Manstein, D. J., Titus, M. A., de Lozanne, A. and Spudich, J. A. (1989). Gene replacement in Dictyostelium: generation of myosin null mutants. EMBO J. 8, 923-932.[Abstract]

Maselli, A., Laevsky, G. and Knecht, D. (2002). Kinetics of binding, uptake and degradation of live fluorescent (DsRed) bacteria by Dictyostelium discoideum. Microbiology 148, 413-420.[Abstract/Free Full Text]

Nellen, W., Datta, S., Reymond, C., Sivertsen, A., Mann, S., Crowley, T. and Firtel, R. A. (1987). Molecular biology in Dictyostelium: tools and applications. Methods Cell Biol. 28, 67-100.[Medline]

O'Halloran, T. J. and Spudich, J. A. (1990). Genetically engineered truncated myosin in Dictyostelium: The carboxyl-terminal regulatory domain is not required for the full development cycle. Proc. Natl. Acad. Sci. USA 87, 8110-8114.[Abstract]

O'Halloran, T. J., Ravid, S. and Spudich, J. A. (1990). Expression of Dictyostelium myosin tail segments in Escherichia coli: domains required for assembly and phosphorylation. J. Cell Biol. 110, 63-70.[Abstract]

Ostrow, B. D., Chen, P. and Chisholm, R. L. (1994). Expression of a myosin regulatory light chain phosphorylation site mutant complements the cytokinesis and developmental defects of Dictyostelium RMLC null cells. J. Cell Biol. 127, 1945-1955.[Abstract]

Parent, C. A., Blacklock, B. J., Froehlich, W. M., Murphy, D. B. and Devreotes, P. N. (1998). G protein signaling events are activated at the leading edge of chemotactic cells. Cell 95, 81-91.[Medline]

Pasternak, C. L., Flicker, P. F., Ravid, S. and Spudich, J. A. (1989a). Intermolecular versus intramolecular interactions of Dictyostelium myosin: possible regulation by heavy chain phosphorylation. J. Cell Biol. 109, 203-210.[Abstract]

Pasternak, C. L., Spudich, J. A. and Elson, E. L. (1989b). Capping of surface receptors and concomitant cortical tension are generated by conventional myosin. Nature 341, 549-551.[CrossRef][Medline]

Peters, D. J. M., Knecht, D. A., Loomis, W. F., de Lozanne, A., Spudich, J. A. and van Haastert, P. J. M. (1988). Signal transduction, chemotaxis, and cell aggregation in Dictyostelium discoideum cells without myosin heavy chain. Dev. Biol. 28, 153-163.

Peterson, M. D., Novak, K. D., Reedy, M. C., Ruman, J. I. and Titus, M. A. (1995). Molecular genetic analysis of myoC, a Dictyostelium myosin I. J. Cell Sci. 108, 1093-1103.[Abstract/Free Full Text]

Pollard, T. D. (1982). Structure and polymerization of Acanthamoeba myosin-II filaments. J. Cell Biol. 95, 816-825.[Abstract]

Robinson, D. N., Cavet, G., Warrick, H. M. and Spudich, J. A. (2002). Quantitation of the distribution and flux of myosin-II during cytokinesis. BMC Cell Biol. 3, 4.[CrossRef][Medline]

Ruppel, K. M., Uyeda, T. Q. P. and Spudich, J. A. (1994). Role of highly conserved lysine 130 of myosin motor domain. In vivo and in vitro characterization of site specifically mutated myosin. J. Biol. Chem. 269, 18773-18780.[Abstract/Free Full Text]

Sellers, J. R. (1999). Myosins. Oxford: Oxford University Press.

Shu, S., Lee, R. J., LeBlanc-Straceski, J. M. and Uyeda, T. Q. P. (1999). Role of myosin II tail sequences in its function and localization at the cleavage furrow of Dictyostelium. J. Cell Sci. 112, 2195-2201.[Abstract/Free Full Text]

Springer, M. L., Patterson, B. and Spudich, J. A. (1994). Stage-specific requirement for myosin II during Dictyostelium development. Development 120, 2651-2660.[Abstract/Free Full Text]

Sussman, S. (1987). Cultivation and synchronous morphogenesis of Dictyostelium under controlled experimental conditions. Methods Cell Biol. 28, 9-29.[Medline]

Thompson, J. D., Gibson, T. J., Plewniak, F., Jeanmougin, F. and Higgins, D. G. (1997). The CLUSTAL-X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 25, 4876-4882.[Abstract/Free Full Text]

Titus, M. A. (1999). A class VII unconventional myosin is required for phagocytosis. Curr. Biol. 9, 1297-1303.[CrossRef][Medline]

Tuxworth, R. I., Weber, I., Wessels, D., Addicks, G. C., Soll, D. R., Gerisch, G. and Titus, M. A. (2001). A role for myosin VII in dynamic cell adhesion. Curr. Biol. 11, 318-329.[CrossRef][Medline]

Uyeda, T. Q. P. and Spudich, J. A. (1993). A functional recombinant myosin II lacking a regulatory light chain-binding site. Science 262, 1867-1870.[Medline]

Uyeda, T. Q. P., Abramson, P. D. and Spudich, J. A. (1996). The neck region of the myosin motor domain acts as a lever arm to generate movement. Proc. Natl. Acad. Sci. USA 93, 14124-14129.

Vailancourt, J. P., Lyons, C. and Côté, G. P. (1988). Identification of two phosphorylated threonines in the tail region of Dictyostelium myosin II. J. Biol. Chem. 263, 10082-10087.[Abstract/Free Full Text]

Vogel, G., Thilo, I., Schwarz, H. and Steinhart, R. (1980). Mechanism of phagocytosis is mediated by different recognition sites as disclosed by mutants with altered phagocytic properties. J. Cell Biol. 86, 456-465.[Abstract/Free Full Text]

Warrick, H., de Lozanne, A., Leinwand, L. and Spudich, J. A. (1986). Conserved protein domains in a myosin heavy chain gene from Dictyostelium discoideum. Proc. Natl. Acad. Sci. USA 83, 9433-9437.[Abstract]

Wessels, D., Soll, D. R., Knecht, D., Loomis, W. F., de Lozanne, A. and Spudich, J. A. (1988). Cell motility and chemotaxis in Dictyostelium amebae lacking myosin heavy chain. Dev. Biol. 128, 164-177.[Medline]

Yumura, S. and Uyeda, T. Q. P. (1997a). Transport of myosin II to the equatorial region without its own motor activity in mitotic Dictyostelium cells. Mol. Biol. Cell 8, 2089-2099.[Abstract/Free Full Text]

Yumura, S. and Uyeda, T. Q. P. (1997b). Myosin II can be localized to the cleavage furrow and to the posterior region of Dictyostelium without control by phosphorylation of myosin heavy and light chains. Cell Motil. Cytoskel. 36, 313-322.[CrossRef][Medline]

Zang, J.-H. and Spudich, J. A. (1998). Myosin II localization during cytokinesis occurs by a mechanism that does not require its motor domain. Proc. Natl. Acad. Sci. USA 95, 13652-13657.[Abstract/Free Full Text]


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