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Genes to Cells (2007) 12, 535-546. doi:10.1111/j.1365-2443.2007.01071.x
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The PI3K-Akt pathway promotes microtubule stabilization in migrating fibroblasts

 Keisuke Onishi1, Maiko Higuchi1, Tomoko Asakura1, Norihisa Masuyama1 and Yukiko Gotoh1,2,*

1 Institute of Molecular and Cellular Biosciences, University of Tokyo, 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-0032, Japan
2 SORST Research Project, Japan Science and Technology Corporation, Tokyo, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
Directed cell migration is controlled by extracellular cues such as growth factors/chemokines and extracellular matrix. In a migrating cell, a subset of microtubules becomes stabilized, and this stabilization is implicated in the establishment and maintenance of cell polarity. It is still not fully understood, however, how extracellular cues regulate the dynamics of microtubules. Here we show that the PI3K-Akt signaling pathway plays a pivotal role in growth factor regulation of microtubule stability. Treatment of NIH 3T3 fibroblasts with platelet-derived growth factor (PDGF) increases the amount of stabilized microtubules, and this increase is abrogated by the addition of a PI3K inhibitor or by expression of a dominant-negative form of Akt (DN-Akt), but not by the addition of a MEK inhibitor. Expression of an active form of Akt slightly increases the bulk amount of stabilized microtubules. Stabilization of microtubules induced in edge cells in the wounded monolayer culture is also attenuated by the PI3K inhibitor treatment or by expression of DN-Akt. Given that Akt is activated at the leading edge of a migrating cell and plays an essential role in directed cell migration, these results reveal a novel mechanism linking extracellular cues to directed cell migration, namely Akt regulation of microtubule stability.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
Directed cell migration plays a crucial role in tissue development, repair and tumor metastasis, and is controlled by extracellular cues, including various soluble factors (cf. growth factors, chemokines) and the extracelullar matrix. Cell migration is initiated by cell polarization, accompanying asymmetric distribution of signaling molecules and asymmetric cytoskeletal organization within the cell. Microtubule dynamics are essential for cell polarization and migration in many cell types such as fibroblasts (Vasiliev et al. 1970) and endothelial cells (Gotlieb et al. 1983) as well as in the nerve growth cones (Tanaka et al. 1995). Perturbation of microtubule dynamics by the addition of low doses of the microtubule-destabilizing agent nocodazole or by the microtubule-stabilizing agent taxol (Liao et al. 1995; Grigoriev et al. 1999) abrogates cell polarity and reduces cell migration into a wound in monolayer cultures. The regulation of microtubule dynamics is thus the key to understanding the molecular basis of directed cell migration.

Microtubule network itself is polarized in migrating cells. In fibroblasts and endothelial cells, a large subset of microtubules are oriented toward the leading edge, and in epithelial cells, microtubule plus ends close to the leading edge were found to grow more persistently than those further back in the cell body (Waterman-Storer & Salmon 1997). Whereas most microtubules exist transiently with a half-life averaging only 5–10 min in cultured fibroblasts (Schulze & Kirschner 1986), a small population of microtubules is more stable than the bulk of microtubules (Schulze & Kirschner 1987). These stabilized microtubules are selectively formed near the leading edge of migrating (wound-edge) cells (Gundersen & Bulinski 1988). Stabilized microtubules are enriched with post-translationally modified tubulins such as acetylated and detyrosinated tubulin (Westermann & Weber 2003). Detyrosinated tubulin (a.k.a. Glu-tubulin), in which the C-terminal tyrosine of {alpha}-tubulin is removed by tubulin carboxypeptidase and the second C-terminal residue glutamate is exposed as the new C-terminus, is proposed to be a consequence rather than a cause of microtubule stability, but this modification may regulate cell migration given that kinesin interacts preferentially with Glu-tubulin in vitro (Liao & Gundersen 1998) and that kinesin-dependent processes, including the recycling of endocytosed transferrin (Lin et al. 2002) and the polarized distribution of vimentin intermediate filaments in migrating cells, depend on stable Glu-tubulin-containing microtubules (Glu microtubules; Gurland & Gundersen 1995). Stabilization of microtubules at the leading edge is also implicated in activation of Rac and actin polymerization, leading to lamellipodial protrusion in migrating cells (Waterman-Storer et al. 1999). Continuous flow of various cargos along microtubules supports extension of the leading edge (Bergmann et al. 1983; Wacker et al. 1997; Hirschberg et al. 1998; Lippincott-Schwartz et al. 2000; Schmoranzer et al. 2003).

So what are the mechanisms that regulate the stability of microtubules in response to extracellular stimuli? Lysophosphatidic acid (LPA) in serum has been shown to induce polarized stable microtubule formation in wound-edge 3T3 fibroblasts through activation of the small GTPase Rho and its effector mDia, which in turn recruits an EB1-APC complex that associates with the plus end of microtubules and stabilizes them, as detected by the increase of Glu microtubules (Wen et al. 2004). Integrin-mediated activation of focal adhesion kinase (FAK) is also involved in the Rho-mDia-mediated microtubule stabilization at the leading edge of 3T3 fibroblasts (Palazzo et al. 2004). On the other hand, in wound-edge CHO and LLCPK1 fibroblasts, inhibition of Rho with C3 transferase had no detectable effect on microtubule dynamics in the leading edge, but stimulated microtubule turnover in the trailing edge, although these cells did exhibit an asymmetric microtubule distribution with more microtubules extending toward the leading edge as a result of selective stabilization of microtubules in this direction (Salaycik et al. 2005). This suggests that mechanisms other than Rho-mediated regulation of microtubules might also contribute to the stabilization of microtubules at the leading edge.

Whereas LPA transmits signals through G protein-coupled receptors, many growth factors that can induce directed cell migration, such as platelet-derived growth factor (PDGF), transmit signals through receptor tyrosine kinases. It has remained unclear whether this class of chemoattractants also induce stabilization of microtubules, and if so, how they do it. A number of signaling molecules, including phosphatidylinositide 3-kinase (PI3K), the small GTPases Rac and Cdc42 and the serine/threonine kinases Akt and ERK, have been shown to mediate growth factor stimulation of cell migration (Higuchi et al. 2001; Matsubayashi et al. 2004). Akt was first described as a proto-oncogene similar to protein kinase A and C. It plays pivotal roles in many physiological contexts such as cell survival, proliferation and metabolism, as well as in pathological contexts such as tumorigenesis and metastasis. Akt is activated downstream of PI3K in response to a variety of growth factors and integrin-mediated signals. Akt becomes active by phosphorylation of two residues T308 and S473, catalyzed by PDK1 and a protein complex called TORC2, respectively, which are regulated downstream of PI3K. Active (phosphorylated) Akt is localized at the leading edge and plays an essential role in promoting cell migration in response to growth factors such as PDGF (Higuchi et al. 2001). It has also been shown that activation of Rac, as well as inactivation of the tumor suppressor gene pten, promotes migration of fibroblasts through activation of Akt (Higuchi et al. 2001). This might in part explain the invasiveness of malignant tumors encompassing with high Akt activities. Some Akt targets such as ACAP1 (Li et al. 2005) and Girdin/APE (Enomoto et al. 2005) have been implicated in the regulation of cell migration. However, the mechanisms by which Akt regulates cell migration have still largely remained elusive, and the relationship between Akt and microtubule stability is yet unknown.

In this study, we find that PDGF stimulation increases the amount of stabilized microtubules in NIH 3T3 fibroblasts and that this increase is mediated by the PI3K-Akt signaling pathway. We also report that the PI3K-Akt signaling pathway is involved in the stabilization of microtubules oriented toward the leading edge of wound-edge cells. These results suggest that, in addition to Rho-dependent mechanism, the PDK1-Akt pathway contributes to the regulation of microtubule stability in response to chemotactic cues at the leading edge.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
Identification of the extracellular cues and signaling molecules that are involved in stabilizing microtubules in the leading edge is a key issue to understanding the regulation of directed cell migration. Although LPA, which activates a G-protein-coupled receptor, has been shown to promote microtubule stabilization, it is unknown whether peptidyl growth factors that activate receptor tyrosine kinases, another class of chemoattractants, can also promote microtubule stabilization. We therefore examined whether PDGF can affect microtubule stability in NIH 3T3 fibroblasts, since PDGF stimulates motility of these cells. We measured the amount of microtubules by cell lysate fractionation, which allows the separation of soluble from cytoskeletal (insoluble) fractions of tubulin (see Experimental procedures). Treatment of NIH 3T3 cells with PDGF reproducibly increased the amount of insoluble tubulin detected by this method at 5 min after PDGF treatment, whereas the total amount of tubulin was unaffected by the PDGF treatment (Fig. 1B). As expected, treatment with the microtubule-destabilizing agent nocodazole abolished the insoluble fraction of tubulin, whereas that with the microtubule-stabilizing agent taxol increased the insoluble and decreased the soluble fraction of tubulin (Fig. 1A), verifying the method used here to detect the amount of polymerized/insoluble tubulin. These data suggest that signaling pathways activated by PDGF treatment can trigger microtubule stabilization within the migrating cell.


Figure 1
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  		Figure 1  PDGF stimulation induces microtubule stabilization. (A) Fractionation and separation of soluble (monomer) and insoluble (polymer) tubulins by centrifugation. NIH 3T3 cells were treated with DMSO vehicle control, nocodazole or taxol for 5 h then harvested in a microtubule stabilization buffer and centrifuged as described in Experimental procedures. The soluble and insoluble fractions were subjected to immunoblotting using an antibody against {alpha}-tubulin. Note that nocodazole treatment reduced the concentration of insoluble tubulin, whereas taxol treatment increased it. (B) Serum-starved NIH 3T3 cells were stimulated with or without 20 ng/mL PDGF for 5 min or 15 min before harvesting. The cell lysates were separated into soluble and insoluble fractions and subjected to immunoblotting with antibody against {alpha}-tubulin. Essentially the same results were obtained in three independent experiments. The normalized insoluble {alpha}-tubulin levels against the total {alpha}-tubulin levels are indicated in arbitrary units.

 
We then analyzed to find which signaling molecules might be involved in the increase in polymerized/insoluble tubulin induced by PDGF treatment. Although the small GTPase Rho is involved in LPA-induced microtubule stabilization, we could not detect activation of Rho in response to PDGF treatment in NIH 3T3 cells (data not shown). Since the PI3K-Akt and MEK-ERK pathways are implicated in growth factor-stimulated motility of fibroblasts, we examined possible roles for these pathways in microtubule polymerization. Pretreatment of NIH 3T3 cells with the PI3K inhibitor LY294002 attenuated the phosphorylation of Akt, but not that of ERK MAP kinase, in response to PDGF treatment (Fig. 2A). Under this condition, LY294002 treatment markedly inhibited the increase of insoluble tubulin induced by PDGF treatment (Fig. 2A). We also found that expression of a dominant-negative form of Akt (DN-Akt) blocked the increase of insoluble tubulin in response to PDGF treatment (Fig. 2B) and that expression of a constitutively active form of Akt slightly, but reproducibly, increased the amount of insoluble tubulin (Fig. 2C). These results suggest that the PI3K-Akt pathway is involved in PDGF-stimulated increase of polymerized/insoluble tubulin, that is, microtubules, in NIH 3T3 cells. On the other hand, pretreatment of the MEK inhibitor U0126 did not affect the level of insoluble tubulin while attenuating the PDGF-induced phosphorylation of ERK (Fig. 2A), suggesting that the MEK-ERK pathway is dispensable for the PDGF-stimulated increase of microtubules.


Figure 2
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  		Figure 2  The PI3K-Akt signaling pathway is necessary for PDGF-induced microtubule stabilization. (A) Serum-starved NIH 3T3 cells were treated with DMSO vehicle control, the PI3K inhibitor LY294002 (20 µM) or the MEK inhibitor U0126 (10 µM) for 1 h then stimulated with or without 20 ng/mL PDGF for 5 min before harvesting. The cell lysates were separated into soluble and insoluble fractions and subjected to immunoblotting with antibodies against {alpha}-tubulin, Akt, phosphorylated Akt (T308) (pAkt), MAPK and phosphorylated MAPK (pMAPK). (B, C) NIH 3T3 cells were transfected with either a control vector or a vector expressing DN-Akt (B) or active Akt (C) for 24 h. In (B), the cells were starved then stimulated with or without 20 ng/mL PDGF for 15 min before harvesting. The cell lysates were separated into soluble and insoluble fractions and subjected to immunoblotting with antibodies against {alpha}-tubulin. Essentially the same results were obtained in three independent experiments. The normalized insoluble {alpha}-tubulin levels against the total {alpha}-tubulin levels are indicated in arbitrary units.

 
Since the local stabilization of microtubules at the leading edge and the role of stabilized microtubules in directed cell migration have been well-established in the wound-healing assay of monolayer cultures, we next asked whether the PI3K-Akt pathway is also involved in the microtubule stabilization in this setting. To detect stabilized microtubules, we first took advantage of antibodies able to specifically recognize post-translationally detyrosinated tubulins (or Glu tubulin), a widely accepted marker of microtubule stability (Schultze et al. 1987; Palazzo et al. 2004). Right after wounding, no obvious accumulation of Glu microtubules was observed, but after 1 h of wounding, Glu microtubules were formed selectively near the leading edge of a subpopulation of wound-edge cells, as previously reported (Gunderson et al. 1988). The overall distribution of total microtubules detected by antibodies to {alpha}-tubulin was indistinguishable between Glu microtubule-positive and -negative cells (Fig. 3A). The tangled and bundled array of Glu microtubules polarized toward the leading edge became more evident after 4 h of wounding (Fig. 3A). Importantly, we found that pretreatment of the monolayer cultures of NIH 3T3 cells with the PI3K inhibitor LY294002 potently inhibited the emergence of Glu microtubules in the wound-edge cells (21.8% ± 2.7% in control vs. 6.9% ± 1.3% in LY294002-treated wound-edge cells; P < 0.01) (Fig. 3B,C). LY294002 treatment preferentially reduced the tangles and bundles of Glu microtubule arrays near the leading edge compared to those near the nucleus. These results suggest that PI3K activity is essential for the microtubule stabilization at the leading edge of the wound-edge cells. In contrast, the MEK inhibitor U0126 had little effect on the formation and array of Glu microtubules in the wound-edge cells (Fig. 3B,C), again suggesting that the MEK pathway is dispensable for microtubule stabilization.


Figure 3
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  		Figure 3  PI3K activity is required for wound-induced Glu-microtubule formation. (A) Distribution of Glu-microtubules in wounded NIH 3T3 cells. NIH 3T3 cells were grown to confluency and then wounded for the indicated times. The cells were then fixed and stained with antibodies against Glu-tubulin and {alpha}-tubulin. Wound edge is toward the top in each panel. (B) The PI3K inhibitor LY294002 but not the MEK inhibitor U0126 suppressed Glu-microtubule formation. NIH 3T3 cells were grown to confluency and treated with DMSO vehicle control, LY294002 (20 µM) or U0126 (10 µM) for 1 h before wounding. After 4 h of wounding, the cells were fixed and stained with antibodies against Glu-tubulin and {alpha}-tubulin. (C) Quantification of ratio of wounded NIH 3T3 cells with Glu-microtubules. Data are the mean ± SEM of values from six samples and similar results were obtained in two independent experiments. *P < 0.01 vs. control; t-test. Scale bar: 50 µm.

 
In addition to Glu microtubules, we also evaluated the extent of microtubule stabilization by detecting nocodazole-resistant (and dilution-resistant) microtubules. In this assay (which is a combination of nocodazole-resistance assay and tubulin-dilution assay; Gundersen & Bulinski 1988), cells are treated with nocodazole for a short period (5 min) in order to selectively depolymerize dynamic microtubules, permeabilized briefly with Triton X-100 to dilute out depolymerized tubulin and then fixed and stained with antibodies to {alpha}-tubulin to visualize microtubules. This assay allowed us to examine the levels of stable microtubules within each cell. Immediately after wounding, nocodazole-resistant microtubules were observed in a sparse and scattered manner in both wound-edge cells and cells away from the wound (Fig. 4A), just like in the confluent cells without wounding (data not shown). However, 1 h (data not shown) or 4 h (Fig. 4A) after wounding, dense arrays of nocodazole-resistant microtubules were formed in almost all the wound-edge cells, but not in the cells away from the wound (Fig. 4A), as reported previously (Gundersen & Bulinski 1988). These nocodazole-resistant microtubules were localized selectively in the region between the nucleus and the leading edge, confirming the polarized formation of stabilized microtubules (Gundersen & Bulinski 1988). Only a subpopulation of nocodazole-resistant microtubules were double-stained with antibodies to Glu tubulin (Fig. 4B), indicating that Glu microtubules are a specific subset of stable microtubules (consistent with a previous report; Gundersen et al. 1987). Using this assay, we asked which signaling molecules were required for microtubule stabilization in the wound-edge cells. Treatment with LY294002 resulted in a dramatic change in the organization of nocodazole-resistant microtubules (Fig. 4C), increasing the proportion of cells without aligned arrays of nocodazole-resistant microtubules from 23.4% ± 2.8% to 55.4% ± 7.2% (P < 0.0005) (Fig. 4D). In addition, existing microtubules were shorter and fewer in number and they were no longer aligned towards the leading edge (Fig. 4C). Furthermore, centrosome-associated microtubules were preferentially lost in LY294002-treated cells. These results demonstrate a pivotal role of PI3K in the regulation of microtubule stability in the wound-healing assay. By contrast, treatment with U0126 had little effects on the array of nocodazole-resistant microtubules (Fig. 4C,D), again suggesting that MEK does not play a major role in the stabilization of microtubules, although previous results have shown that it transmits the wound signal to orient centrosome against the wound in MDCK cells (Matsubayashi et al. 2004).


Figure 4
Figure 4
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  		Figure 4  PI3K activity is required for wound-induced formation of nocodazole-resistant microtubules. (A) Distribution of nocodazole-resistant microtubules in wounded NIH 3T3 cells. NIH 3T3 cells were grown to confluency and wounded for the indicated times then treated with 5 µM nocodazole for 5 min. The cells were permeabilized with a PEM buffer containing 0.2% TX-100 to visualize nocodazole-resistant microtubules, then fixed and stained with antibodies against {alpha}-tubulin and ß-catenin to visualize cell-cell boundary and nucleus. Wound edge is toward the bottom in each panel. (B) Nocodazole-resistant microtubules overlapped with Glu-tubulin. NIH 3T3 cells treated as in (A) were stained with antibodies against Glu-tubulin and {alpha}-tubulin. Note that Glu-microtubules were a subset of nocodazole-resistant microtubules formed in wound-edge cells. (C) PI3K inhibitor suppressed and GSK3 inhibitor enhanced wound-induced nocodazole-resistant microtubule formation. NIH 3T3 cells were grown to confluency and treated with DMSO vehicle control, LY294002 (20 µM), U0126 (10 µM), rapamycin (20 nM) or SB216763 (25 µM) for 1 h before wounding. After 4 h of wounding, the cells were treated with 5 µM nocodazole for 5 min, permeabilized, fixed and stained with antibodies against {alpha}-tubulin and ß-catenin as in (A). (D) Quantification of ratio of wounded NIH 3T3 cells with nocodazole-resistant-microtubules in (C). We determined the percentage of cells along the wound edge that do not contain microtubules that were aligned and oriented towards the leading edge (polarized microtubules). Data are the mean ± SEM of values from six samples and similar results were obtained in two independent experiments. *P < 0.0005 vs. control; t-test. Scale bar: 50 µm.

 
We then asked which of the signaling molecules activated by PI3K were involved in the regulation of nocodazole-resistant microtubules. Whereas transfection of a control vector did not affect the array of nocodazole-resistant microtubules, transfection of a vector expressing a DN-Akt markedly suppressed the formation of nocodazole-resistant microtubules at the leading edge of the wound-edge cells (29.3% ± 1.9% in control vs. 52.2% ± 1.3% in DN-Akt expressing wound-edge cells, P < 0.005) (Fig. 5B). These results provide further evidence that Akt is a key downstream effector of PI3K in the regulation of microtubule stability. We next examined the possible involvement of the kinases mammalian target of rapamycin (mTOR), which is activated by the PI3K-Akt signaling pathway and glycogen synthase kinase 3 (GSK3), which is inhibited by the PI3K-Akt pathway, since these kinases have been implicated in the regulation of cell motility (Berven et al. 2004; Eng et al. 2006). Inhibition of the mTOR/raptor complex by treatment with rapamycin did not suppress the formation of nocodazole-resistant microtubules significantly, although the array of these microtubules in rapamycin-treated cells appeared slightly affected compared to that in control cells (Fig. 4C). On the other hand, inhibition of GSK3 by SB216763 increased the amount of nocodazole-resistant microtubules. Notably, SB216763 treatment stimulated the formation of nocodazole-resistant microtubules not just in the wound-edge cells but also in the cells away from the wound and resulted in a loss of polarization of these microtubules in cells located at the wound. This suggests that GSK3 might be important for suppressing the stabilization of microtubules in regions other than the leading edge of the wound-edge cells. This is consistent with the notion that Akt promotes microtubule stabilization through inhibition of GSK3, although it is also possible that Akt and GSK3 regulate microtubule stability independently (see Discussion).


Figure 5
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  		Figure 5  Akt activity is necessary for wound-induced formation of nocodazole-resistant microtubules. (A) NIH 3T3 cells were transfected with either a control vector or a vector expressing DN-Akt for 24 h. Histone H2B-GFP was used as a transfection marker. The confluent cells were wounded for 4 h then treated with 5 µM nocodazole for 5 min, permeabilized, fixed and stained with antibodies against {alpha}-tubulin and ß-catenin. (B) Quantification of ratio of wounded NIH 3T3 cells with nocodazole-resistant-microtubules in (A). We determined the percentage of cells along the wound edge that do not contain microtubules that were aligned and oriented towards the leading edge (polarized microtubules). Data are the mean ± SEM of values from four samples and similar results were obtained in two independent experiments. *P < 0.01 vs. control; t-test. DN-Akt inhibited wound-induced nocodazole-resistant microtubule formation. Scale bar: 50 µm.

 

   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
The plus ends of microtubules alternate between growing and shrinking phases, a process called "dynamic instability", and explore intracellular spaces to search for cues that capture the plus ends and stabilize them. This "search-and-capture" of microtubules was proposed about two decades ago to explain the generation of asymmetric microtubule arrays and polarization of the cell (Kirschner & Mitchison 1986). In migrating fibroblasts, a subset of microtubules ("pioneer microtubules") oriented toward the leading edge indeed becomes stabilized due to a decrease in catastrophe frequency (Wittmann et al. 2003). The prevailing hypothesis is that these stabilized microtubules serve as tracks for the directed delivery of membrane vesicles to the leading edge, contributing to the establishment and maintenance of the front-back cell polarity and promoting the cell's forward advance. It is therefore important to clarify how a cell decides when and where stabilized microtubules are formed, to understand the regulation of directed cell migration. In this study, we have revealed that the PI3K-Akt pathway plays a pivotal role in the stabilization of microtubules by the use of three different assays. First, the amount of polymerized tubulin detected by cell lysate fractionation was increased in response to PDGF treatment, and this increase was inhibited by the blockade of either PI3K or Akt activity. Moreover, expression of active Akt slightly increased the amount of polymerized tubulin in this assay. Second, selective formation of Glu microtubules in the subset of wound-edge cells in the wound-healing assay was suppressed by the inhibition of PI3K activity. Third, selective formation of nocodazole-resistant microtubules at the leading edge of the wound-edge cells was also suppressed by the blockade of either PI3K or Akt activity. Since the PI3K-Akt pathway is activated in response to various chemoattractants such as PDGF, epidermal growth factor, fibroblast growth factor, hepatocyte growth factor, vascular endothelial growth factor and neurotrophins, as well as to integrin-mediated extracellular matrix signals, this pathway might commonly contribute to the spatio-temporal regulation of microtubule stability. In fact, the activity of the PI3K-Akt pathway is highly localized to the very front of the leading edge (Higuchi et al. 2001). It is therefore likely that the PI3K-Akt pathway serves as a key determinant of the spatial restriction of stabilized microtubules to the leading edge, as a part of the "capture" mechanism.

Active Rac has been shown to generate pioneer microtubules within the leading edge (Wittmann et al. 2003). We found that Akt promotes microtubule stabilization in the leading edge, and since Akt is activated downstream of Rac, Akt might be involved in mediating the Rac regulation of microtubule stability. This might explain at least in part why Akt is necessary for Rac promotion of cell motility (Higuchi et al. 2001).

We found that both PI3K and Akt play roles in microtubule stabilization. However, Akt might not be the only mediator of PI3K regulation of microtubule stability. For example, a recent paper showed that LL5ß, a PH domain containing protein, is localized to the cell cortex by binding to the PI3K product PIP3, and recruits CLASP, a microtubule plus end tracking protein (+TIPS), together with its accessory protein ELKS (Lansbergen et al. 2006). The observations suggest that there are parallel pathways operating downstream of PI3K that orient microtubule arrays to the leading edge.

Our results show that inhibition of GSK3, but not of mTOR, robustly affects microtubule stability. Although GSK3 is a well-known target of Akt, this does not necessarily mean that Akt regulates microtubule stability through inhibition of GSK3, especially given that the activity of GSK3 can be regulated by other molecules such as aPKC and Wnt signaling. In fact, a recent paper utilizing the GSK3{alpha}/ß double knockin mouse has shown that Akt phosphorylation sites of GSK3{alpha} and ß are dispensable for GSK3 regulation of growth cone motility, a microtubule-dependent process considered to be very similar to fibroblast motility (Gartner et al. 2006). Therefore, it is still an open question what target molecule(s) is responsible for Akt regulation of microtubule stability. Several mechanisms have been proposed to anchor +TIPs to the cell cortex, including recruitment of the LL5ß-ELKS-CLASP complex (mentioned above), ACF7-CLASP (Drabek et al. 2006), Cdc42/Rac-IQGAP-CLIP170 (Fukata et al. 2002) and Rho-mDia-EB1/APC (Wen et al. 2004), where CLASP, CLIP170, EB1 and APC are +TIPs. It will be interesting to determine whether Akt participates in the regulation of these mechanisms.

Even in the absence of chemoattractant cues, a cell often becomes polarized spontaneously and forms a single leading edge in the front. A positive feedback loop of phosphatidylinositol 3,4,5-trisphosphate (PIP3), where PIP3 recruits more PIP3 locally within the cell membrane, has been suggested to contribute to such a spontaneous cell polarization as well as to chemoattractant-triggered cell polarization (Wang et al. 2002; Weiner et al. 2002). The positive feedback loop of PIP3 that takes within minutes does not need microtubules, and in fact some cell types including neutrophils can establish cell polarity and migrate without microtubule arrays (Niggli 2003). In contrast, other cell types that are considered to require stable/sustained cell polarity to migrate long distances including fibroblasts (Vasiliev et al. 1970) and endothelial cells (Gotlieb et al. 1983), as well as the growth cones of neuronal cells (Tanaka et al. 1995), require microtubules to establish polarity and migration. It is conceivable that the latter system (microtubule-dependent, sustained front-back polarity) is established based on the former system (microtubule-independent, transient front-back polarity). If this is the case, the PIP3-Akt-mediated stabilization of microtubules described in this study might be a good candidate to link between these two systems.


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Plasmids, reagents and antibodies

Active Akt [PH domain (residues 4–129)-truncated Akt with a myristoylation site at its N-terminus] and dominant-negative Akt (DN-Akt, K179A) were kindly provided by Drs R. Roth and D. Alessi, respectively. Dominant-negative Akt was amplified by PCR and inserted into the expression vector pCS2+ (Masuyama et al. 2001). We obtained PDGF from R&D, LY294002, U0126 and rapamycin from Calbiochem, SB216763 from TOCRIS and nocodazole from Sigma. The antibodies used in this study include anti-Akt (Cell Signaling), anti-phospho-Akt (Thr308) (Santa Cruz), anti-ERK1 (K-23, Santa Cruz Biotechnology), anti-phospho-MAPK (Promega), anti-{alpha}-tubulin (DM1{alpha}, Sigma) for immunoblotting, anti-detyrosinated (Glu)-tubulin (Chemicon), anti-ß-catenin (Sigma), Alexa 488-conjugated anti-mouse IgG antibody and Alexa 594-conjugated anti-rabbit IgG antibody (Molecular Probes) for cell staining.

Cell lines and transfection

NIH 3T3 cells were maintained in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum. Cell transfection was carried out using Lipofect AMINE PLUS reagent (Invitrogen). For six-well plates, NIH 3T3 cells were plated on coverslips at a density of 1 x 105/well and incubated for 48 h. Cells were then transfected with 1 µg of total DNA together with 6 µL of PLUS reagent and 4 µL of LipofectAMINE reagent/well. For detecting transfected cells in cell staining experiments, histone H2B-GFP was used as a transfection marker.

Nocodazole-resistance assay and cell staining

NIH 3T3 cells were plated on coverslips and cultured for 72 h. After the indicated times of wounding, the cells were incubated with 5 µM nocodazole for 5 min. At this point, cells were incubated for 1 min at 37 °C with 0.2% Triton X-100 in PEM buffer (100 mM PIPES [pH 6.9], 1 mM EGTA, 2 mM MgCl2) to remove monomeric tubulin, then rinsed twice in PEM buffer and finally fixed in methanol 10 min at –20 °C. The cells were permeabilized with 0.2% Triton X-100 in phosphate-buffered saline (PBS) for 5 min and incubated with 5% fetal bovine serum in PBS for 30 min to block nonspecific antibody binding. {alpha}-tubulin, ß-catenin and Glu-tubulin (detyrosinated-tubulin) were detected using a mouse anti-{alpha}-tubulin monoclonal antibody, a rabbit anti-ß-catenin polyclonal antibody and a rabbit anti-detyrosinated-tubulin polyclonal antibody, respectively. After washing with PBS, the cells were stained with Alexa 488-conjugated anti-mouse IgG antibody or Alexa 594-conjugated anti-rabbit IgG.

Measurement of soluble or insoluble tubulin and immunoblotting

NIH 3T3 cells transfected with plasmids or pretreated with pharmacological inhibitors as described were lyzed in Microtubule Stabilization Buffer (MSB) (85 mM PIPES [pH 6.9], 1 mM EGTA, 1 mM MgCl2, 2 M Glycerol, 0.5% Triton X-100) after the indicated PDGF stimulation times. Lysates were kept 2 min at 4 °C and then centrifuged 5 min at 15 300 g Supernatants, representing the soluble fraction of proteins, were transferred to new tubes and 5 x Laemli buffer was added. Pellets, representing the polymerized fraction of proteins, were washed once in MSB then resuspended in 2 x Laemli buffer. Cell lysates were subjected to immunoblotting with anti-Akt, anti-phospho-Akt (Thr308), anti-MAPK, anti-phospho-MAPK and anti-{alpha}-tubulin antibodies. Intensities of insoluble and total {alpha}-tuulin bands were measured with IMAGEJ software (Rasband, WS, ImageJ, U.S. National Institutes of Health, Bethesda, MD, (< http://rsb.info.nih.gov/ij/>).


   Acknowledgements
 
We thank Drs Dario R. Alessi and Richard Roth for Akt constructs and Dr Elizabeth Nigh for critical reading of the manuscript. We also thank members of the Gotoh laboratory for encouragement and helpful discussions. This work was supported by Grants-in-aid from the Ministry of Education, Culture, Sports, Science and Technology of Japan as well as SORST of Japan Science and Technology Corporation.


   Footnotes
 
Communicated by: Eisuke Nishida

* Correspondence: E-mail: ygotoh{at}iam.u-tokyo.ac.jp


   References
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
 References
 
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Received: 12 January 2007
Accepted: 20 January 2007




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