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Genes to Cells (2009) 14, 821-834. doi:10.1111/j.1365-2443.2009.01312.x
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PTEN is a mechanosensing signal transducer for myosin II localization in Dictyostelium cells

 Md. Kamruzzaman Pramanik1,2, Miho Iijima3, Yoshiaki Iwadate1,4 and Shigehiko Yumura1,*

1 Department of Functional Molecular Biology, Graduate School of Medicine, Yamaguchi University, Yamaguchi 753-8512, Japan
2 Radioisotope Production Division, INST, AERE, Bangladesh Atomic Energy Commission, Bangladesh
3 Johns Hopkins University, School of Medicine, Baltimore, MD 21205, USA
4 PRESTO, Japan Science and Technology Agency, 4-1-8, Honcho Kawaguchi, Saitama 332-0012, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
To investigate the role of PTEN in regulation of cortical motile activity, especially in myosin II localization, eGFP–PTEN and mRFP–myosin II were simultaneously expressed in Dictyostelium cells. PTEN and myosin II co-localized at the posterior of migrating cells and furrow region of dividing cells. In suspension culture, PTEN knockout (pten–) cells became multinucleated, and myosin II significantly decreased in amount at the furrow. During pseudopod retraction and cell aspiration by microcapillary, PTEN accumulated at the tips of pseudopods and aspirated lobes prior to the accumulation of myosin II. In pten– cells, only a small amount of myosin II accumulated at the retracting pseudopods and aspirated cell lobes. PTEN accumulated at the retracting pseudopods and aspirated lobes even in myosin II null cells and latrunculin B-treated cells though in reduced amounts, indicating that PTEN accumulates partially depending on myosin II and cortical actin. Accumulation of PTEN prior to myosin II suggests that PTEN is an upstream component in signaling pathway to localize myosin II, possibly with mechanosensing signaling loop where actomyosin-driven contraction further augments accumulation of PTEN and myosin II by a positive feedback mechanism.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
PTEN (phosphatase and tensin homolog deleted on chromosome 10), also known as MMAC1, is a tumor suppressor protein that regulates a variety of biological processes and suppresses tumor formation, primarily by its lipid phosphatase activity (Steck et al. 1997; Myers et al. 1998; Li et al. 2005). Mutation or deletion of PTEN is commonly found in a wide range of human tumors, such as endometrial and prostate carcinomas, breast cancer and glioblastomas. Thus, PTEN is one of the most frequently mutated tumor suppressors (Li et al. 1997). PTEN dephosphorylates phosphatidylinositol 3,4,5-triphosphate (PIP3) into phosphatidylinositol 4,5-bisphosphate (PIP2) and influences PI3 kinase-Akt signaling, which regulates multiple downstream signaling mechanisms, including cell proliferation and apoptosis (Maehama & Dixon 1998; Leslie & Downes 2002). It has been demonstrated that PTEN also modulates cell migration (Gu et al. 1999).

Recent studies have indicated that PTEN regulates directional migration of neutrophils and Dictyostelium during chemotaxis (Iijima & Devreotes 2002; Li et al. 2005; Billadeau 2008). Dictyostelium discoideum has proven to be an excellent model system to study the molecular mechanisms of chemotaxis and cell polarity signals because these mechanisms are highly conserved from higher eukaryotes to this social amoeba. In chemotaxing Dictyostelium cells, PI3 kinase (PI3K) localizes at the leading edge and PTEN, a negative regulator of the PI3K pathways, localizes at the lateral and posterior regions of migrating cells (Funamoto et al. 2002). This reciprocal localization of PI3K and PTEN confines PIP3 production at the leading edge of the cell and thus mediates directional actin polymerization, pseudopod formation, and chemotaxis. PTEN knockout or knockin cells show defective directional migration or chemotaxis (Funamoto et al. 2002; Iijima & Devreotes 2002). In chemotaxing neutrophils, a similar pattern of PTEN localization has been reported (Li et al. 2003, 2005; Wu et al. 2004). Knockdown of PTEN through siRNA inhibits the chemotaxis of Jurkat cells (Li et al. 2005).

Similar to PTEN, myosin II is localized at the posterior region of both migrating neutrophils and Dictyostelium cells, which confers posterior contraction (Yumura et al. 1984; Uchida et al. 2002; Parent 2004). During cell division, PTEN and myosin II localize at the furrow region in Dictyostelium cells (Yumura 1996; Janetopoulos et al. 2005). In myosin II heavy chain null cells or myosin II heavy chain phosphorylation mutants, Dictyostelium cells produce multiple pseudopods and show impaired chemotaxis (Spudich 1989; Heid et al. 2004). Similar defects are also observed in pten– cells (Iijima & Devreotes 2002; Wessels et al. 2007). In shaking culture, both pten– and myosin II null cells become multinucleated, albeit to a lesser extent in pten– cells (De Lozanne & Spudich 1987; Janetopoulos et al. 2005).

As both of these proteins are localized to the same sites and confer many common behavioral defects when defective, we speculated that PTEN may play an important role in the localization of myosin II during cell migration and other cellular processes, including cytokinesis. Recently, it has been suggested that PTEN plays a role in suppressing lateral pseudopod formation during chemotaxis (Wessels et al. 2007); however, there is no strong evidence to solidify the relationship between these two proteins in cell motility and cytokinesis.

Mechanosensation, the sensing capability of mechanical forces, is a fundamental cellular process that allows cells to sense their surrounding environment and respond accordingly. Mechanosensation has been implicated in the performance of diverse functions, such as tissue formation, blood-pressure regulation, muscle contraction, hearing, bone remodeling, and cell shape control (Kee & Robinson 2008). Mechanosensing and mechanical feedback are also considered fundamental for cell shape regulation during cytokinesis (Effler et al. 2007). There are three main classes of proposed mechanosensors: entire actin networks, stretch-activated membrane channels, and motor proteins (Martinac 2004; Tamada et al. 2004; Effler et al. 2006; Orr et al. 2006). Myosin II has been proposed as a candidate for providing mechanosensory function (Effler et al. 2007; Kee & Robinson 2008).

In the present study, we examined the role of PTEN in myosin II localization, using Dictyostelium cells as a model organism. By co-expressing both proteins tagged with eGFP and mRFP, we observed their co-localization over a time course during migration and cytokinesis. We also found that accumulation of PTEN preceded that of myosin II, with a difference of several seconds at the retracting pseudopod as well as at a part of the cell aspirated by microcapillary. Furthermore, myosin II did not properly accumulate at the subcellular regions of pten– cells. Therefore, we concluded that PTEN is an upstream signal transducer for myosin II localization. Moreover, in myosin II null cells and latrunculin B-treated wild-type cells, PTEN responded to the mechanical distortion of the cell membrane and localized at the tip of aspirated cell lobes, though the localization of PTEN decreased in both cases compared with the control cells. Therefore, PTEN can respond to mechanical stimulation independently of myosin II and actin filaments but the accumulation of PTEN is augmented by interaction between these two cytoskeletal proteins. Here we propose that PTEN contributes to the subcellular localization of myosin II through mechanosensation and the actomyosin-driven contraction furnishes a positive feedback signal to this mechanosensing signaling loop.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Co-localization of PTEN and myosin II during cell migration and cytokinesis

To analyze the spatial and temporal correlation between both PTEN and myosin II, Dictyostelium cells were simultaneously transformed with extrachromosomal plasmids containing eGFP–PTEN (hereafter GFP–PTEN) and mRFP–myosin II (hereafter, RFP–myosin II) genes. The localizations of two proteins were simultaneously observed during cell migration and cytokinesis under a confocal microscope. At the onset of cytokinesis, the cells rounded up, and both GFP–PTEN and RFP–myosin II were intensely localized over the entire cortex. As the cytokinesis progressed, both proteins accumulated at the equatorial region (Fig. 1A). Co-localization of GFP–PTEN and RFP–myosin II were also observed at the posterior and lateral regions of the migrating cells (Fig. 1B). Sometimes, accumulation of both proteins was found at the anterior pseudopods and, in many cases, such pseudopods tended to retract as described later.


Figure 1
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  		Figure 1  Co-localization of PTEN and myosin II in cytokinetic and migrating cells. Dictyostelium cells simultaneously expressing GFP-PTEN and RFP-myosin II were observed under a confocal microscope. (A) GFP-PTEN and RFP-myosin II were co-localized at the furrow region of the dividing cells. (B) Both proteins were also co-localized at the posterior region of migrating cells. Arrow shows the direction of cell migration. Bar, 10 µm.

 
Aberrant localization of myosin II in dividing pten– cells

To investigate the role of PTEN in the function of myosin II, pten– cells were newly generated by homologous recombination, wherein the majority of the PTEN gene, including the catalytic domain, was deleted. We generated 5 clones of pten– cells and ascertained that all of the clones showed similar phenotypes. The growth rate of the pten– cells was similar to that of wild-type (AX2) cells when grown on substratum. The generation times of pten– and wild-type cells were 13.93 and 11.94 h, respectively (Fig. S1A in Supporting Information/Supplementary material). In shaking culture, growth rate of pten– cells was significantly lower than that of wild-type cells, and generation times of pten– and wild-type cells were 18.35 and 12.64 h, respectively (Fig. S1B). Fig. S1C,D shows DAPI staining of both cell types, indicating that pten– cells became multinucleated in shaking culture. Approximately 60% of pten– cells were multinucleated, whereas this was the case for only about 20% of wild-type cells (Fig. S1E,F). Therefore, PTEN seems to play an important role in cytokinesis under suspension condition.

To clarify the reason for this, wild-type and pten– cells expressing GFP–myosin II were embedded in 0.03% low-melting temperature agarose to mimic the suspension condition, and GFP–myosin II was observed during cytokinesis under a confocal microscope. In wild-type cells, GFP–myosin II localized at the furrow (Fig. 2A), similar to that of cells attached to substratum (Fig. 7A). Figure 2(B) shows fluorescence intensity profiles demonstrating that the highest fluorescence peak was found at the constricting furrow region (Fig. 2B). The fluorescence ratio, measured as a ratio of fluorescence intensity at the furrow and the cytoplasm outside the furrow region, increased as the cell division proceeded, and reached 1.97 ± 0.34 (averaged ratio ± SD, n = 6) during active constriction of the furrow ring (Fig. 2E,F). However, GFP–myosin II scarcely localized at the furrow region of dividing pten– cells (Fig. 2C,D). The maximum fluorescence ratio between the furrow region and the cytoplasm was only 1.11 ± 0.07 (n = 5, Fig. 2G,H). Therefore, lack of PTEN induced aberrant localization of myosin II under this condition.


Figure 2
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  		Figure 2  Aberrant localization of myosin II at the furrow region in pten– cells during cytokinesis. Wild-type and pten– cells expressing GFP-myosin II were embedded in 0.03% low-melting agarose to mimic the suspension condition and observed under a confocal microscope. (A) In wild-type cells, myosin II localized at the furrow region during cell division. Bar, 10 µm. (B) Fluorescence intensity profiles across the cell during cytokinesis as marked by rectangular selection in panel A. (C) In pten– cells, little amount of myosin II localized at the furrow region. Bar, 10 µm. (D) Fluorescence intensity profiles across the cell as marked by rectangular selection in panel C. (E, F) Time course of fluorescence ratio of myosin II localization at the furrow region (ratio of fluorescence intensity at the furrow and cytoplasm) of dividing wild-type cell, (indicated as rectangles in panel F). (G, H) Time course of fluorescence ratio of myosin II localization at the furrow region of dividing pten– cell, (indicated as rectangles in panel H). To observe individual myosin II filaments during cytokinesis mimicking suspension condition, cells expressing GFP–myosin II were treated with 5 mM EDTA, which detached the cells from the substratum. After slightly compressed with agarose sheet, the cells were observed under a TIRF microscope. (I) A substantial amount of myosin II filaments localized at the equatorial region in wild-type cells. (J) In pten– cells, initially myosin filaments localized at the equatorial region but eventually disappeared from the furrow and localized at the polar region of one daughter cell. Disappearance of myosin II filaments ceased furrow ingression leading to failure in cytokinesis. White arrows indicate the location of nuclei in the dividing cell. Bar, 10 µm.

 

Figure 7
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  		Figure 7  Involvement of PTEN in mechanosensing signaling under agar-overlay condition. (A) Without agar-overlay, the amount of myosin II at the furrow region of pten– cells was less than wild-type cells. (B) Under agar-overlay, the amount of myosin II at the furrow region increased in wild-type cells with a pressure of agar-overlay. In the case of pten– cells, myosin II increased slightly. (C) Bar diagrams show the fluorescence ratios (fluorescence intensity at furrow and cytoplasm) of wild-type and pten– cells without agar-overlay (white bar) and under agar-overlay (gray bar) condition. Note that the fluorescence ratios at furrow region in wild-type and pten– cells are significantly different without agar-overlay and agar-overlay as depicted by P-values (a = 1.12 x  10–10 and b = 4.81 x 10–7, respectively). P-values were considered significant if P < 0.05 (Student's t-test, assuming unequal variances). Bars, 10 µm.

 
To examine how individual myosin II filaments accumulate at the furrow region in pten– cells in suspension conditions, cells expressing GFP–myosin II were observed in the presence of 5 mM EDTA (ethylenediamine tetraacetic acid), which detached the cells from the substratum. The cells were slightly pressed with a thin agarose sheet to keep their position (Yumura et al. 1984), which enabled us to observe individual filaments under a total internal reflection fluorescence (TIRF) microscope. In the case of wild-type cells, myosin II filaments were localized at the equatorial region, and all of the observed cells completed cell division (Fig. 2I). On the contrary, about 24% of pten– cells (5 out of 21) failed to complete cytokinesis under this condition. Initially, myosin II filaments localized at the equatorial region to a lesser extent than in wild-type cells. In the cases of failed cytokinesis, myosin II filaments eventually began to disappear from the equatorial region and appear at the either polar region (Fig. 2J). Disappearance of myosin II filaments from the furrow region ceased furrow ingression, leading to failure in cytokinesis. The density of myosin II filaments in the cortex of pten– cells was lower than that of wild-type cells, suggesting that PTEN may contribute to the stable binding of myosin filaments to the cortex.

Sequential localization of PTEN and myosin II in retracting pseudopods

Previously, it was reported that multiple pseudopods were frequently observed in pten– cells, as well as in myosin II null cells (Spudich 1989; Iijima & Devreotes 2002; Wessels et al. 2007). In migrating cells, myosin II accumulates at the tip of the extended pseudopods and contributes to the subsequent retraction of pseudopods (Moores et al. 1996; Yumura 1996). Thus, PTEN may contribute to pseudopod retraction by localizing myosin II. To examine this possibility, GFP–PTEN and RFP–myosin II were simultaneously observed in retracting pseudopods under a confocal microscope. RFP–myosin II localized at the tip of retracting pseudopods, as reported previously (Fig. 3A). Interestingly, GFP–PTEN was also localized at the retracting pseudopods about 13.5 ± 9.2 s (n = 11) earlier than RFP–myosin II (Fig. 3B). Figure 3(C) shows kymographs of sequential localization of GFP–PTEN and RFP–myosin II, indicating that PTEN localization preceded that of myosin II before pseudopod retraction. When myosin II began to localize, the pseudopod initiated retraction (arrows in Fig. 3B).


Figure 3
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  		Figure 3  GFP-PTEN and RFP-myosin II co-localize sequentially at the retracting pseudopods. Dictyostelium cells simultaneously expressing GFP–PTEN and RFP–myosin II were observed under a confocal microscope. (A) GFP–PTEN and RFP–myosin II localization in retracting pseudopod of migrating cell. During extension, no substantial localization of GFP–PTEN and RFP–myosin II was observed at the pseudopod (0 s). Before beginning of retraction, PTEN appeared at the tip of pseudopod (42 s). After several seconds, myosin II followed PTEN's localization at the same site (63 s). White arrowheads indicate the localization sites of GFP–PTEN and RFP–myosin II at retracting pseudopod. Bar, 10 µm (B) Time course of fluorescence intensity of GFP-PTEN (circles), RFP-myosin II (squares) and the length of pseudopod (triangles). White and black arrows indicate starting points of localization of GFP–PTEN and RFP–myosin II, respectively. (C) Kymographs of GFP–PTEN and RFP–myosin II fluorescence of the rectangle depicted in panel A.

 
Next, we examined whether the myosin II properly accumulates at the tip of retracting pseudopods in pten– cells. Only a small amount of GFP–myosin II accumulated at the tip of retracting pseudopods of pten– cells, contrary to the results in wild-type cells (Fig. 4A,B). Accumulation of myosin II was also reduced at the posterior region of migrating pten– cells (Fig. S2). During pseudopod extension, the fluorescence ratio between the tip of pseudopod and cytoplasm was almost constant in both wild-type and pten– cells. As the pseudopod was retracting, the fluorescence ratio gradually increased and reached 3.88 ± 0.84 (n = 15) in wild-type cells (Fig. 4C,E). In contrast, the maximum fluorescence ratio in pten– cells was only 1.44 ± 0.41 (n = 8) during the retraction phase (Fig. 4D,F).


Figure 4
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  		Figure 4  Little amount of myosin II localized at the retracting pseudopod of pten– cells. Retracting pseudopods of wild-type and pten– cells expressing GFP–myosin II were observed under a confocal microscope. (A) Time-lapse images of a retracting pseudopod showing GFP–myosin II in wild-type cell. Bar, 10 µm. (B) Time-lapse images of a retracting pseudopod showing GFP–myosin II in pten– cell. Bar, 10 µm. (C, D) Kymographs of GFP–myosin II in pseudopod of wild-type and pten– cells made from the rectangles depicted in panels A and B, respectively. (E, F) Time course of fluorescence ratio of GFP–myosin II at the tip of the pseudopod (circles) and changes in the length of pseudopod (triangles) shown in panels A and B, respectively. (G) Retraction speed of pseudopods in wild-type, pten– and myosin II null cells. Note that the retraction speed is significantly slow in pten– and myosin II null cells as depicted by different P-values (a = 3.4 x 10–6, b = 2.3 x 10–7 and c = 0.16). Asterisks indicate statistically significant difference compared with wild-type cells. P-values were considered significant if P < 0.05 (Student's t-test, assuming unequal variances).

 
We also measured the relative speed of pseudopod retraction in wild-type, pten–, and myosin II null cells (Fig. 4G). The relative speeds of pseudopod retraction in pten– cells and myosin II null cells were similar (0.18 ± 0.05 µm/s, n = 48 and 0.16 ± 0.05 µm/s, n = 31, respectively) but significantly (evaluated by Student's t-test) slower than that of wild-type cells (0.25 ± 0.09 µm/s, n = 48). Taken together, the accumulation of PTEN as well as myosin II at the pseudopods contributes to their retraction. The accumulation of PTEN prior to myosin II suggests that PTEN is in the upstream of signaling pathway to localize myosin II.

PTEN and myosin II localize sequentially, responding to the mechanical distortion of cell cortex

When part of a Dictyostelium cell is aspirated by a microcapillary, GFP–myosin II accumulates at the tip of aspirated cell lobe, indicating that myosin II responds to the mechanical distortion (Merkel et al. 2000). Merkel and colleagues found these mechanosensing responses of myosin II in interphase cells. Subsequently, Effler et al. described that GFP–myosin II localization by aspiration occurs only in mitotic cells, not in interphase cells (Effler et al. 2007). To resolve this contradiction and to examine whether PTEN also localizes in response to the mechanical load, aspiration experiments were carried out with the cells simultaneously expressing GFP–PTEN and RFP–myosin II. When part of the interphase cells was aspirated by a microcapillary, both of these proteins localized at the tip of aspirated lobes sequentially (Fig. 5A). Similar responses were also observed in mitotic cells (data not shown), suggesting that mechanosensing property exists both in mitotic and interphase cells. Quantitative analysis on kymographs clearly showed that GFP–PTEN was accumulated at the tip about 8.87 ± 0.85 s (n = 5) earlier than RFP–myosin II (Fig. 5B,C). When RFP–myosin II began to accumulate, the aspirated lobe began to retract. These experiments indicate that PTEN is an upstream component in the mechanosensing signaling pathway for myosin II localization and retraction of the cell cortex.


Figure 5
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  		Figure 5  Co-localization of both GFP–PTEN and RFP–myosin II in response to mechanical distortion of cell cortex. Part of a Dictyostelium cell simultaneously expressing GFP–PTEN and RFP–myosin II was aspirated with a microcapillary and the localizations of these two proteins were observed under a confocal microscope. (A) Time lapse images of GFP–PTEN and RFP–myosin II in response to the mechanical distortion of cortex. White arrowheads indicate the localization sites of GFP–PTEN and RFP–myosin II in the aspirated lobe. Bar, 10 µm. (B) Time course of fluorescence intensity of GFP–PTEN (circles) and RFP–myosin II (squares) and changes in the length of aspirated lobe (triangles). GFP–PTEN localization preceded that of RFP–myosin II. Pseudopod began to retract at the same time when myosin II began to accumulate at the tip. White and black arrows indicate starting points of localization of GFP–PTEN and RFP–myosin II, respectively. Bar, 10 µm. (C) Kymographs of GFP–PTEN and RFP–myosin II fluorescence of the rectangle depicted in panel A.

 
To assess this possibility, aspiration experiments were carried out using pten– cells expressing GFP–myosin II. A significantly reduced amount of GFP–myosin II was observed at the tip of aspirated lobe of pten– cells compared to wild-type cells (Fig. 6A,B). The accumulation pattern of GFP–myosin II at the aspirated tips of pten– and wild-type cells are shown in the kymographs (Fig. 6C,D). Maximum fluorescence ratio in wild-type cells (measured as the ratio of fluorescence intensity at the aspirated tip and cytoplasm) was 3.18 ± 0.73 (n = 5), whereas it was only 1.65 ± 0.37 (n = 8) in the case of pten– cells (Fig. 6E,F).


Figure 6
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  		Figure 6  Less amount of myosin II accumulates in response to the mechanical distortion of cortex in pten– cells. (A) Time-lapse images of an aspirated lobe showing GFP–myosin II localization in wild-type cell. Bar, 10 µm (B) Time-lapse images of an aspirated lobe in pten– cell. Note that significantly reduced amount of myosin II accumulated at the tip of aspirated lobe in pten– cell. Bar, 10 µm (C, D) Kymographs of GFP–myosin II in aspirated lobs of wild-type and pten– cells made by the rectangles depicted in panels A and B, respectively. (E, F) Time course of fluorescence ratio (circles) of GFP–myosin II at the tip and changes in the length of aspirated lobe (triangles) in wild-type (E) and pten– cells (F). Note that pten– cells took longer time to retract the aspirated lobe than wild-type cells. Bar, 10 µm.

 
Reduced amount of myosin II localized at the furrow region of dividing pten– cells under compression with agar-overlay

Previously, Yumura and Uyeda reported that when dividing cells are flattened by an agarose sheet, the amount of myosin II increased at the furrow region (Yumura & Uyeda 2003). Cells compressed with an agarose sheet may transduce the same mechanical signal involving PTEN, as in the case of retracting pseudopods and aspirated cell lobes. To investigate this possibility, we compared the extent of myosin II localization at the furrow region of cytokinetic wild-type and pten– cells when adhered to glass surface with or without compression by agar-overlay. In pten– cells, reduced amount of myosin II localized under both conditions (Fig. 7A,B). Without agar-overlay, fluorescence ratio of GFP–myosin II at the furrow region of wild-type and pten– cells were 2.23 ± 0.37 (average ratio ± SD, n = 14) and 1.53 ± 0.24 (average ratio ± SD, n = 19), respectively (Fig. 7C). Under agar-overlay, fluorescence ratios of GFP–myosin II at the furrow region of wild-type and pten– were 3.89 ± 0.98 (average ratio ± SD, n = 29) and 2.22 ± 0.31 (average ratio ± SD, n = 14), respectively. Thus the amount of myosin II localized at the furrow in pten– cells was reduced compared with the wild-type when compressed, suggesting that PTEN partially contributes to the increased accumulation of myosin II under agar-overlay condition through the mechanosensing signaling pathway.

Actomyosin contraction provides a positive feedback signal for PTEN localization

Previous studies showed that PTEN localizes in response to chemotactic stimulation independently of actin filaments (Iijima et al. 2004; Hoeller & Kay 2007). To examine the contribution of actin filaments in localization of PTEN in response to the mechanical signal, cells expressing GFP–PTEN were treated with 7.5 µM latrunculin B for 20 min. This concentration of the drug completely disrupted the actin cytoskeleton (data not shown). Following the latrunculin B treatment, cell aspiration experiments were carried out as described above. The latrunculin B-treated cells still showed localization of GFP–PTEN at the tip of aspirated lobes, although the quantity was decreased (Fig. 8A,B). Kymographs in Fig. 8(D,E) show the PTEN localization along with time course. Cells could not retract the lobes in the presence of latrunculin B, and consequently, the length of the lobes gradually increased. Average peaks of fluorescence ratio in control and latrunculin B-treated cells were 3.09 ± 0.88 (n = 5) and 1.82 ± 0.45 (n = 5), respectively (Fig. 8G,H). Therefore, PTEN is able to accumulate independently of cortical actin filaments at the tip of lobes.


Figure 8
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  		Figure 8  PTEN can localize in response to mechanical distortion independently of actin and myosin II. Cells expressing GFP–PTEN were treated with 7.5 µM latrunculin B for 30 min. (A) Time lapse images of GFP–PTEN during aspiration with a microcapillary in the absence of latrunculin B. (B) Time lapse images of GFP–PTEN in the presence of latrunculin B. (C) Time lapse images of GFP–PTEN in myosin II null cell. Note that less amount of GFP–PTEN accumulated at the tip of latrunculin B-treated and myosin II null cells than untreated wild-type cells. (D, E, F) Kymographs of GFP–PTEN localization at the aspirated cell lobes of control (untreated), latrunculin B-treated and myosin II null cells, respectively. (G, H, I) Time courses of fluorescence ratio (circles) of GFP–PTEN at the tip and length of aspirated lobe (triangles) are shown in panels A, B and C in the absence and presence of latrunculin B and myosin II null cells, respectively. Note that the latrunculin B-treated and myosin II null cells could not retract the lobes. Bars, 10 µm.

 
Although PTEN appears to be an upstream signal for myosin II localization as shown above, the accumulated myosin II at the tip of aspirated lobes generates force, which may, in turn, lead to more PTEN localization and, consequently, more myosin II localization. To examine whether myosin II contributes to PTEN localization, myosin II null cells expressing GFP–PTEN were aspirated by a microcapillary. A reduced amount of GFP–PTEN was localized at the tip of aspirated lobes (Fig. 8C). The kymograph in Fig. 8(F) shows the PTEN localization along with time course. As myosin II null cells could not contract the lobes, length of the lobes gradually increased. Figure 8(I) shows that the peak of fluorescence ratio of GFP–PTEN was only 1.4 ± 0.17 (n = 5), compared with that of wild-type cells (3.09 ± 0.88, n = 5). Thus, PTEN is able to accumulate at the aspirated cortices independently of actin and myosin II, but these cytoskeletal proteins enhance further accumulation of PTEN possibly by a positive feedback mechanism.


   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
During chemotaxis, PTEN localizes at the posterior and lateral sides of the cells (Funamoto et al. 2002; Iijima & Devreotes 2002). In the present study, we found that PTEN also tends to localize at the posterior region in unstimulated, spontaneously polarized cells, though stimulated cells show much clearer accumulation. This observation is reasonable, as PTEN localizes at the furrow region of a dividing cell, which is not stimulated but can be considered as two well-polarized, nascent cells migrating in opposite directions. The PTEN localization pattern is similar to that of myosin II in both neutrophils and Dictyostelium. We have demonstrated co-localization of these two proteins during migration and cytokinesis in Dictyostelium cells simultaneously expressing GFP–PTEN and RFP–myosin II. During pseudopod retraction, GFP–PTEN localized several seconds earlier than RFP–myosin II at the tip of the pseudopods. Pseudopod retraction occurred following the myosin II localization. On occasion, exact co-localization did not take place during migration, which can be explained by the sequential localization. When PTEN localized at one place, myosin II localization initiated at the same site, but if the cell changes the polarity rapidly, myosin II and PTEN localization may not coincide exactly, as there is time difference in localizing these two proteins. Sequential co-localization of GFP–PTEN and RFP–myosin II was also observed at the distorted cell membrane by aspiration. Therefore, PTEN plays an important role in subcellular localization of myosin II.

The pten– cells were multinucleated in shaking culture, like myosin null cells, though not to the equivalent extent. Multinucleation was not severe in pten– cells when adhered to the substratum. It is likely that pten– cells employ attachment-assisted or traction mediated-cytokinesis (cytokinesis B), in which cells exert opposite pulling forces from both daughters and can complete cell division (Uyeda et al. 2004). In this case, myosin II-mediated contraction is dispensable. Conversely, in shaking culture, furrow ingression is absolutely dependent on myosin II-mediated contraction, and if myosin II localization is perturbed, it is plausible that the cells will consequently fail to perform cytokinesis. A reduced amount of myosin II was localized at the furrow region of pten– cells while embedded in agarose, mimicking suspension conditions. Therefore, myosin II localizes at the furrow region of the cytokinetic cell primarily in a PTEN-dependent manner. Under TIRF microscopy, fewer myosin II filaments accumulated at the cell periphery at the onset of cytokinesis in pten– cells, and they began to delocalize from the equatorial region. This mislocalization of myosin II broke the cytokinetic cell symmetry which finally led cells to give up cytokinesis. Therefore, PTEN not only helps recruitment of myosin II at the equatorial region but also contributes to the persistent localization of myosin II filaments at the furrow region.

In the present study, we found that PTEN accumulates at the tip of retracting pseudopods. Furthermore, a reduced amount of myosin II localized at the retracting pseudopods of pten– cells compared to wild-type cells. The speed of pseudopod retraction was also lower in pten– cells in comparison with that of wild-type cells, which was expected, as pseudopod retraction is contributed by accumulated myosin II (Yumura 1996). Similarly, in the cell aspiration experiment, less GFP-myosin II was accumulated at the aspirated tip of pten– cells than wild-type cells. All these observations indicate that myosin II localization is, at least partially, PTEN-dependent.

Mechanosensing and mechanotransduction are fundamental to a wide variety of cellular processes such as hearing, bone remodeling, blood pressure regulation, and exercise-induced skeletal muscle growth. Cytoskeletal proteins and stretch-activated channels have been implicated in different mechanosensing processes (Tamada et al. 2004; Orr et al. 2006). Effler et al. proposed that myosin II is a candidate for providing mechanosensory functions during cytokinesis in Dictyostelium (Effler et al. 2007). Myosin II exhibits mechanosensing property because ADP-release from myosin II slows when strain is applied (Kee & Robinson 2008). The mechanosensory response is also important for monitoring the cell shape during cytokinesis, because mechanical perturbation can lead to the defective cytokinesis by causing disturbance in mechanosensory system (Effler et al. 2006). Cortical deformation by micropipette aspiration also induces mechanical strain on the cortical actin network, which could be sensed directly by myosin II. However, in the present study, PTEN localized at the retracting pseudopods or at the aspirated cell lobes earlier than myosin II, and only a reduced amount of myosin II accumulated in pten– cells. These observations suggest that PTEN is an upstream regulator of such mechanosensing signals for myosin II localization. To retract a pseudopod or aspirated cell lobe, cells recruit PTEN and myosin II, which ultimately generate a retraction force. This retraction force may give positive feedback to the mechanosensory system, because both PTEN and myosin II localization gradually increased after initiating the pseudopod retraction (Fig. 3).

Myosin II localization still occurred, although less prominently, in retracting pseudopods and aspirated lobes even in the absence of PTEN. Multiple signaling pathways for myosin II localization have been proposed (Bosgraaf & Haastert 2006). Myosin II translocation to the cortex is considered to occur by two steps: first, myosin II self-assembles into bipolar filaments in the cytosol. Filament formation is negatively regulated by myosin II heavy chain phosphorylation, through the action of several myosin heavy chain kinases (MHCK A, B C, D), and positively regulated by dephosphorylation of the heavy chains by phosphatase PP2A (Murphy & Egelhoff 1999; Yumura et al. 2005). Secondly, the association of filaments to the cortex occurs in an actin-dependent manner. This association is regulated by the cGMP-binding protein GbpC, and possibly PAKa and RegA; PAKa, in turn, is regulated by Rac and Akt/PKB (Chung et al. 2001; Bosgraaf et al. 2002; Bosgraaf & Haastert 2006). The accumulation of myosin II was also proposed to be mediated by increased concentrations of Ca2+ and calmodulin, which were shown to inhibit the myosin heavy chain kinase that causes filament disassembly (Yumura & Kitanishi-Yumura 1993). Once the filaments are associated with actin, this actomyosin complex can exert contraction force at localized regions, such as posterior regions of migrating cells or furrow regions of dividing cells. This motor activity is regulated by the phosphorylation of the regulatory light chain by myosin light chain kinase A (MLCK A), which is activated by phosphorylation with cGMP-activated kinase GbpC directly or indirectly and subsequent autophosphorylation (Smith et al. 1996; Bosgraaf & Haastert 2006).

At present, no direct connection has been demonstrated between PTEN and myosin II in signaling pathways as far as we know. Beside its lipid phosphatase activity, PTEN has protein phosphatase activity (Myers et al. 1997). Since PTEN has tyrosine phosphatase activity, it is unable to dephosphorylate the threonine phosphates on myosin heavy chains. The assembly of myosin II into bipolar filaments is regulated by the phosphorylation of three threonine residues in the tail region of myosin II molecules (Vaillancourt et al. 1988). Similarly, because the phosphorylation of serine on regulatory light chains of myosin II regulates motor activity, PTEN cannot dephosphorylate it. Therefore, it is not plausible that PTEN directly regulates assembly or motor activity of myosin II via the dephosphorylation of the heavy and light chains. Under TIRF microscopy, PTEN and myosin II were observed as dot-like structures and filaments, respectively (data not shown). They did not show exact co-localization, again suggesting that they did not directly associate with each other in vivo. However, PTEN may contribute to myosin II localization indirectly through its product PIP2 produced from dephosphorylation of PIP3. This PIP2 has been suggested to regulate myosin II localization indirectly by binding and regulating the function of cytoskeletal associated proteins e.g. anillin and ERM in mammalian cells, which in turn contribute to assembly of actomyosin ring at the furrow region (Logan & Mandato 2006). Contractility of actomyosin structure may further be accomplished by PLC-mediated hydrolysis of PIP2 and downstream activation of Ca2+/CaM and PKC.

The major cytoskeletal protein, actin, is essential for myosin II localization. In latrunculin B-treated cells, myosin II does not localize anywhere of the cortex (Yumura et al. 2008). Interestingly, PTEN localized at the tip of aspirated cell lobes in the presence of latrunculin B, indicating that PTEN signaling is independent of actin filaments. However, as the amount of PTEN at the tip of aspirated lobes decreased in latrunculin B-treated cells, actin may partially facilitate or enhance the localization of PTEN by binding the PTEN tensin homolog domain or another unknown mechanism. In myosin II null cells, a smaller amount of PTEN accumulated at the tip of aspirated lobes, indicating that myosin II is a requisite for proper accumulation of PTEN. Since PTEN accumulated earlier than myosin II in retracting pseudopods and aspirated lobes, PTEN may initiate the signaling for myosin II localization, which in turn causes actomyosin contraction and tension. This actomyosin contraction again may cause more PTEN localization and thus input a positive feedback to the mechanosensing signaling loop, wherein PTEN, actin and myosin II contribute together mutually. The identification of mechanosensing molecules and the elucidation of the mechanism of mechanosensing feedback loop are our next challenges.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Cell culture

Dictyostelium discoideum (strain AX2) cells were cultured axenically in HL-5 medium (1.3% bacteriological peptone, 0.75% yeast extract, 85.5 mM D-glucose, 3.5 mM Na2HPO4·12H2O, and 3.5 mM K2HPO4, pH 6.4) at 22 °C. For adherent culture, cells were grown on 9 cm plastic dishes; for shaking cultures, cells were cultured at 22 °C in a reciprocal shaker at 150 rpm. For culturing the cells expressing GFP–PTEN and RFP-myosin II, medium was supplemented with 10 µg/mL geneticin (G418, Sigma-Aldrich), 4 µg/mL blasticidin (Kaken), or both, as required. Before observation, HL-5 medium in plastic dishes was replaced with BSS (10 mM NaCl, 10 mM KCl, 3 mM CaCl2, and 2 mM MES, pH 6.4), and, to obtain actively migrating cells, cells were starved for about 6–7 h.

Co-expression of GFP–PTEN and RFP–myosin II and construction of pten– cells

For simultaneous observation of PTEN and myosin II localization, Dictyostelium cells were first transformed by electroporation with a plasmid DNA construct containing GFP–PTEN gene (Yumura et al. 1995) and transformed colonies were selected by geneticin. Previous experiments, including our earlier experiments, used wild-type PTEN as a source of GFP-fusion protein, but over-expression of the full-length protein dramatically decreased localization of myosin II (data not shown). Probably, excess amount of phosphatase activities of PTEN affects the localization of myosin II. Here, we used PTEN (G129E) as a source of GFP-fusion protein, in which the glycine residue at the position 129 was replaced with glutamate, which previously showed negligible phosphatase activity and same localization as wild-type PTEN (Iijima et al. 2004). G418-resistant transformants were again transformed by electroporation with a plasmid containing the RFP–myosin II gene and transformed colonies were selected by blasticidin. After selection of double resistant colonies, cells were checked for co-expression of both proteins by fluorescence microscopy.

Pten– cells were constructed in the AX2 background. After digesting pBluescript plasmid containing PTEN disruption vector (pPTENdis) with Not1 and Sal1, the linearized pPTENdis was separated by agarose gel electrophoresis and purified by ethanol/acetate precipitation method. Purified linearized pten– construct was introduced in the vegetative AX2 cells by electroporation. Blasticidin was added for selection 12 h after electroporation. After 5 days, blasticidin resistant colonies were isolated and checked for homologous recombination by PCR.

Soft agarose embedding

To mimic the suspension condition, we adopted the soft agarose embedding technique as previously described by Gerald et al. (1998) with slight modification. Fifty microliters of cell suspension in BSS was added to 100 µL of 0.1% melted agarose (Low Melting Temperature Agarose-A9414, Sigma-Aldrich) in a microtube and mixed at 22 °C and mounted in the chamber for observation.

Cell aspiration

To observe the localization of GFP–PTEN and RFP–myosin II upon membrane distortion, cell aspiration technique was adopted. Glass microcapillaries with an inner diameter of 4.0–6.0 µm were prepared by a puller (PG-1, Narishige, Japan) and a micro-forge (MF-830, Narishige, Japan). They were connected to a micrometer-positioned water manometer, by which specific suction pressure can be applied by changing the height of water column. The needle position was manipulated by a micromanipulator (Sigma Koki, Japan) attached to an inverted confocal microscope (LSM-510 META, Zeiss) equipped with a 100x objective (Plan Neofluar, NA 1.3). After positioning the microneedle to the cell, aspiration was carried out continuously by maintaining the suction pressure at a particular level. Time-lapse imaging was carried out simultaneously for analysis.

TIRF microscopy

Dividing cells were observed by TIRF microscopy system of our own composition as previously described (Tokunaga et al. 1997) to study the dynamics of individual myosin II filaments. TIRF microscope was assembled based on an inverted microscope (Olympus, IX71, Japan) with a 60x objective lens (PLAPON60X OTIRFM, NA 1.45, Olympus). A laser diode (DPBL-9010F, Photop Technologies, China) was used for excitation of GFP fused protein at a wavelength of 473 nm. Electric shutters in front of the laser heads controlled exciting light from the laser. The fluorescence of GFP was detected with a dual band-pass filter (XF3056, Omega Optical). Fluorescence images were collected by a digital cooled CCD camera (ORCA ER, Hamamatsu Photonics, Japan). The shutter and CCD camera were controlled using a custom program developed under Lab VIEW 7.1 (National Instruments, Japan). For mimicking suspension conditions, cells were treated with 5 mM EDTA solution (10 mM KCl, 10 mM NaCl, 0.1 mM MgCl2, 5 mM EDTA, 5 mM MES, pH 6.4). To observe myosin II filaments in the ventral cortex under TIRF microscopy, cells were slightly compressed by an agarose sheet containing the same solution (Yumura et al. 1984).

Fluorescence and confocal microscopy

To determine the number of nuclei per cell, cells were stained with 4,6-diamidino-2-phenylindole (DAPI) as previously described (Yumura et al. 2005). After cells were collected by centrifugation at 300 g for 1.5 min, they were fixed in 15 mM phosphate buffer containing 2.5% formaldehyde. An aliquot of cell suspension was mixed with 1 µg/mL of DAPI reagent on a glass slide. The cells were overlaid with a 22 x 22 mm coverslip and mildly pressed with filter paper. Cells were examined by a fluorescence microscope (Nikon, DIAPHOT-300) upon UV illumination with a 40x phase contrast objective.

Cells expressing GFP-PTEN and RFP-myosin II were observed under a confocal microscope (Zeiss, LSM-510 META) equipped with a 100x objective (Plan Neofluar, NA 1.3). Cells were washed with BSS twice and mounted with agar overlay as previously described (Yumura et al. 1984). Time lapse imaging of the specimen was carried out by the confocal microscope using an argon laser at 488 nm for excitation of GFP–PTEN and, simultaneously, a He–Ne laser at 543 nm for excitation of RFP–myosin II. Fluorescence due to GFP and RFP was detected with 505–530 nm band-pass and 560 nm low-pass filter, respectively.

Image analysis

After acquiring the confocal images, co-localization of GFP-PTEN and RFP-myosin II was determined by processing the image-stacks with ImageJ (http://rsb.info.nih.gov/ij/). Kymographs were made from the rectangular selection of the movies over time courses. Fluorescence intensity on the edge-line of retracting region in the kymograph was quantified by a plug-in in ImageJ, which was written by Dr Arai in Osaka University. Fluorescence intensities of GFP and RFP for every image were measured by plot profile option along the time course. Onset of fluorescence accumulation was determined at the rising point of the plot from where the differences between two successive Y-coordinates began to increase consistently. Relative fluorescence ratio at the furrow regions was determined as the ratio of fluorescence intensities at the furrow and cytoplasm.


   Acknowledgements
 
The authors would like to thank Dr Firtel for providing GFP-PTEN in the early work. We also thank Dr Arai and Mr Utsumi for providing the plug-in for ImageJ and image capturing software, respectively. We also thank Drs Uyeda and Kitanishi-Yumura for stimulating discussions and helpful comments. A part of this work was supported by Grants-in-aid for Scientific Research in Priority Areas and Grant-in-Aid for Exploratory Research from the MEXT of Japan.


   Footnotes
 
Communicated by: Masao Tasaka

* Correspondence: yumura{at}yamaguchi-u.ac.jp


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Received: 5 February 2009
Accepted: 13 April 2009




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