|
|
Eukaryot Cell. 2008 June; 7(6): 958–966. Published online 2008 April 11. doi: 10.1128/EC.00442-07. | PMCID: PMC2446661 |
Copyright © 2008, American Society for Microbiology MPL1, a Novel Phosphatase with Leucine-Rich Repeats, Is Essential for Proper ERK2 Phosphorylation and Cell Motility ![[down-pointing small open triangle]](/corehtml/pmc/pmcents/x25BF.gif) Marbelys Rodriguez,† Bohye Kim,† Nam-Sihk Lee, Sudhakar Veeranki, and Leung Kim* Department of Biological Sciences, Florida International University, Miami, Florida 33199 Received December 5, 2007; Accepted April 1, 2008. The novel Dictyostelium phosphatase MPL1 contains six leucine-rich repeats at the amino-terminal end and a phosphatase domain at the carboxyl end. Similarly architectured phosphatases exist among other protozoa, such as Entamoeba histolytica, Leishmania major, and Trypanosoma cruzi. MPL1 was strongly induced after 5 h of development; ablation by homologous recombination led to defective streaming and aggregation during development. In addition, cyclic AMP (cAMP)-pulsed mpl1− cells showed reduced random and directional motility. At the molecular level, mpl1− cells displayed higher prestimulus and persistent poststimulus ERK2 phosphorylation in response to cAMP stimulation. Consistent with their phenotype of persistent ERK2 phosphorylation, mpl1− cells also displayed an aberrant pattern of cAMP production, resembling that of the regA− cells. Reintroduction of a full-length MPL1 into mpl1− cells restored aggregation, ERK2 regulation, random and directional motility, and cAMP production similar to wild-type cells. We propose that MPL1 is a novel phosphatase essential for proper regulation of ERK2 phosphorylation and optimal motility during development. Mitogen-activated protein kinases (MAPKs) are central in the regulation of proliferation, differentiation, and cell migration in diverse eukaryotic cells ( 10, 13, 21, 25, 26, 27). The MAP kinase ERK2 also plays critical roles during Dictyostelium development. ERK2 is essential for initiation and propagation of periodic cyclic AMP (cAMP) pulses during aggregation and differentiation. Chemoattractants, such as cAMP, induce ERK2 activation. Activated ERK2 subsequently inhibits the intracellular cAMP-specific phosphodiesterase RegA, resulting in an increase in the cytosolic cAMP level ( 15, 19, 22). erk2− cells, starved for 8 h, exhibited a decrease in motility and a severe chemotaxis defect toward a cAMP gradient. Aberrancy in chemotaxis was aggravated in the presence of a strong cAMP gradient (2 μM) compared to a weak one (0.1 μM) ( 27). erk2− cells also display defective cytoskeletal remodeling in response to chemoattractant stimulation. A polarized wild-type cell typically displays a single dominant leading edge enriched with F-actin. Myosin II localizes to the lateral side and back of a polarized cell, where it functions to suppress lateral pseudopods and provides tractional force to the back. This single dominant leading edge disintegrates but forms again after 7 min in response to global cAMP stimulation ( 27). In contrast, erk2− cells, under the same condition, displayed multiple crown-like membranous protrusions, which were enriched not only in F-actin but also in myosin II ( 27). This aberrant structure, which was proposed to be less stable and unable to provide necessary traction force for cells to move, is believed to be the reason why erk2− cells are less motile than wild-type cells ( 27). It is well-established that the dual phosphorylation of threonine 176 and tyrosine 178 residues of the ERK2 activation loop, often called the TEY motif, mediates activation of ERK2 kinase activity. This dual phosphorylation on the TEY motif of ERK2 peaks around 1 min after cAMP stimulation, but virtually no phosphorylation of the ERK2 remains after 2 to 3 min ( 14, 27). Adaptation of ERK2 is thus likely mediated by a phosphatase that can dephosphorylate phosphates from both threonine 176 and tyrosine 178 residues. In Dictyostelium, several tyrosine phosphatases (PTP1, PTP2, and PTP3) have been characterized ( 4, 6, 7, 8, 9). These phosphatases are involved in cell differentiation or stress response signaling, but it is not known if ERK2 is regulated by any of these tyrosine phosphatases. Currently, the phosphatase responsible for dephosphorylating ERK2 in Dictyostelium is unidentified. In mammalian systems, several phosphatases are known to decrease MAPK phosphorylation and activity. These MAPK phosphatases belong to the dual specificity phosphatase (DSPase) family, which dephosphorylate both phospho-serine/threonine and phospho-tyrosine residues. Two well-characterized examples of DSPases are Cdc25 and MAP kinase phosphatase (MKP). Cdc25 dephosphorylates and activates cell cycle-dependent kinases to promote cell cycle progression. MKP dephosphorylates and inactivates MAP kinase signaling at the level of MAP kinase ( 2, 18, 20). A number of mammalian MKPs contain a MAP kinase binding (MKB) domain, but no such domain can be found in the Dictyostelium genome by homology domain search (NCBI, Conserved Domains Search BLAST). We have isolated and characterized the function of MPL1, a novel Dictyostelium phosphatase with a leucine-rich repeat (LRR) domain. The MPL1 phosphatase domain contains the conserved, functionally critical signature sequence of DSPases: Dx 26(V/L)x(V/I)HCxAG(I/V)SRSxT(I/V)xxAY(L/I)M (x can be any amino acid). This sequence constitutes a unique structure that enables DSPase to dephosphorylate all three types of phospho-amino acids ( 11, 23). MPL1 contains an LRR domain in its N-terminal region that is composed of six LRR sequences, matching the consensus sequence of LxxLxLxxN/CxL (x can be any amino acid, and L can be leucine, isoleucine, or phenylalanine). The LRR domain often forms a horseshoe-shaped domain capable of mediating diverse protein-protein interactions in many eukaryotes ( 5, 12). The combination of potential DSPases with LRR domains can be found not only in Dictyostelium but also in other parasitic unicellular eukaryotic organisms, such as Entamoeba histolytica, Leishmania major, and Trypanosoma cruzi ( 16) (Fig. ). Ablation of MPL1 resulted in higher prestimulus and persistent poststimulus ERK2 phosphorylation upon cAMP stimulation. Furthermore, mpl1− cells displayed strong defects in motility. Similar to regA− cells, mpl1− cells displayed more rapid cAMP production during the 2-min window after the stimulation with cAMP compared to wild-type cells. Considering that ERK2 is a negative regulator of RegA, persistent activation of ERK2 would have resulted in a persistent RegA inhibition in mpl1− cells. Consistently, reintroduction of the full-length MPL1 in mpl1− cells restored ERK2 regulation, motility, and cAMP production. We propose that MPL1 is a novel phosphatase essential for proper regulation of ERK2 phosphorylation and effective cell movement during Dictyostelium development. Dictyostelium culture and development. Dictyostelium cells were grown in axenic medium (7.15 g peptone 3 [Difco], 7.15 g thiotone E peptone [Becton Dickinson], 7.15 g yeast extract, 15.4 g glucose, 0.525 g Na2HPO4·7H2O, 0.48 g KH2PO4 in 1 liter of water). Cells were developed on nitrocellulose filters at 1 × 107 cells/cm2 in DB buffer (10 mM sodium phosphate, pH 6.4, 2 mM MgCl2, and 0.2 mM CaCl2), or about 100 actively growing Dictyostelium cells were mixed with 200 μl of a saturated overnight culture of Klebsiella aerogenes and plated on a 100 mm SM agar plate (10 g glucose, 10 g Bacto Peptone, 1 g yeast extract, 1 g MgSO4·7H2O, 1.9 g KH2PO4, 0.6 g K2HPO4, 20 g agar in 1 liter of water). Full-length MPL1 cloning. The full-length MPL1 cDNA is 2,505 bp long and contains a single EcoRI site at +405, a SpeI site at +652, and a StyI site at +1742 ( http://dictybase.org; sequence information for DDB0190671). The 662-bp partial MPL1 cDNA fragments encoding the 5′ end of the gene (fragment I, +1 to +662), was generated by PCR using the forward primer 5′-CGGGATCCATGATATTTAAAAAATTATTTTCAAAAGG-3′ and the reverse primer 5′-GGGAAACTAGTGAATTGATTAATAC-3′ and subcloned into pBluescript II KS(−) [pKS(−); Stratagene] vector using BamHI and SpeI sites. The 3′ end of the gene (fragment II, +1742 to +2505), 763 bp in length, was generated by PCR using the forward primer 5′-GAATGCGGCCGCGCCAAGGAAAGATTCAGC-3′ and the reverse primer 5′-CGAGCTCTTATTTTGATAAATCTTTTTCAAATTTTTTTAATTGG-3′ and subcloned into the pKS(−) vector containing fragment I using NotI and SacI sites. The third partial cDNA encoding the central MPL1 cDNA region (fragment III, +663 to +1741) was generated by reverse transcription-PCR (RT-PCR) using the forward primer 5′-GTATTAATCAATTCACTAGTTTCCC-3′ and the reverse primer 5′-GCTGAATCTTTCCTTGGCTTTC-3′. The full-length MPL1 was generated by inserting fragment III into the pKS(−) vector containing the two partial MPL1 genes described above using SpeI and StyI sites. The Flag tag was added to the 5′ end of the full-length MPL1 by PCR using the forward primer 5′-ATAAGCTTTAATAAAAAATGGACTACAAGGACGACGATGACAAGATGATATTTAAAAAATTATTTTCAAAAGG-3′ and the reverse primer 5′-GGAATTTCATAGAATTCCATATAG-3′. The PCR product was digested with HindIII and EcoRI and subcloned into the pKS(−) vector containing full-length MPL1. Then, the construct was digested with XhoI and SacI and subcloned into the pCR 2.1-TOPO vector (Invitrogen) using the same sites. A positive clone was digested with HindIII and subcloned into an empty pKS(−) vector to introduce new restriction sites. Clones having BamHI at the 5′ side and XhoI at the 3′ side were selected, digested with the same enzymes, and subcloned into the pEXP4(+) vector ( 3) previously digested with BclII and XhoI. All PCR products were confirmed by sequencing both strands after each subcloning step. Generation of mpl1− cells and Northern blot analysis of MPL1. The blasticidin resistance cassette was subcloned between the LRR and phosphatase (PPase) domain (Gly 583) of MPL1 (Fig. ). Transformants were prescreened by PCR (with the forward primer 5′-GCATCTGATAATACTGATGAGGC-3′ and the reverse primer 5′-TTCAAGTTGCTGAATCTTTCC-3′). Genomic DNAs from the knockout candidates were isolated, and 5 μg of each sample was digested with EcoRV (see Fig. , below). Knockout cells were analyzed by genomic Southern blotting. The MPL1 expression pattern was determined by Northern blot analysis using a partial MPL1 cDNA encoding the phosphatase domain. The levels of the residual partial MPL1 transcripts in mpl1− cells were compared to that of wild-type cells by RT-PCR using the forward primer 5′-ATGATATTTAAAAAATTATTTTCAAAAGG-3′ and the reverse primer 5′-GGGAAACTAGTGAATTGATTAATAC-3′, resulting in a product that is 662 bp in length upstream of the blasticidin cassette insertion point. One ng of total RNA from either wild-type or mpl1− cells was used for the RT-PCR template for MPL1 amplification. Ig7 transcripts were amplified using the forward primer 5′-GGTGAGCGAAAGCCGAGGAGAG-3′ and the reverse primer 5′-GCAACAGTTACGGGTTCCGCC-3′ as a control. Ten pg of total RNA from either wild-type or mpl1− cells was used for RT-PCR template for Ig7 amplification. Recombinant MPL1 phosphatase and phosphatase assay. The MPL1 phosphatase domain was initially amplified by RT-PCR with the forward primer 5′-CCAAGGAAAGATTCAGCAAC-3′ and the reverse primer 5′-TTATTTTGATAAATCTTTTTCAAATTTTTTTAATTGG-3′ and cloned into the pCR2.1-TOPO vector. Positive clones were selected after sequencing the whole region. GST-MPL1-PPase was generated by subcloning the MPL1 phosphatase domain into pGEX 4T-2 (Pharmacia Inc.) after EcoRI digestion and filling in with Klenow. Clones containing the phosphatase domain were screened first by PCR and confirmed by fusion protein expression of a 53-kDa product upon isopropyl-β-d-thiogalactopyranoside induction. Glutathione S-transferase (GST) and GST-MPL1-PPase proteins were purified with glutathione-Sepharose beads (Pharmacia Inc.) and analyzed by Western blotting using anti-GST antibody. MPL1 phosphatase activities were determined by measuring p-NP generation from p-NPP (Enzolyte pNPP phosphatase assay kit; AnaSpec). The generation of p-NP was determined by monitoring absorption at 405 nm. Submerged aggregation assay. For submerged aggregation experiments, log-phase cells were harvested, washed, and placed under DB buffer at the indicated cell densities on a 24-well plate (Falcon 353047; Becton Dickinson). After 12 h at 20°C, cell migration, streaming, and aggregation were monitored using a Leica inverted microscope (DM IRB). Chemotaxis and random motility analyses. Log-phase cells were differentiated with 50 nM cAMP pulses at 6-min intervals for 4 h. Pulsed cells were plated at a density of 3 × 104 cells/cm2 on a 35-mm tissue culture dish cover (Falcon 353001; Becton Dickinson). A Schmazu micromanipulator with a glass capillary needle (Eppendorf Femtotip) filled with either 0.1 μM or 2 μM cAMP solution was used for a chemotaxis assay. For random motility analysis, cells were plated on the same tissue culture plate cover with no cAMP source, and their movements were recorded for 30 min. The responses of the cells were followed by time-lapse video recording with OpenLab software. Chemotaxing cells were analyzed as described previously ( 17). An ellipsoid was constructed around the cell to allow the same centroid for both the ellipsoid and the contour. The chemotactic index, defined as the net distance moved to the direction of the pipette divided by the total distance moved, was computed from the centroid positions. The speeds of movement of cells were calculated from positions of the centroids. Statistical significance of differences between wild-type and mutant cells were evaluated by obtaining P values using Student's t test. ERK2 activation by cAMP. Both wild-type and mpl1− cells (108 cells) were stimulated with 50 nM cAMP pulses for 4 h, treated with 2 mM caffeine for 30 min, and then stimulated with 0.1 μM cAMP. Cells were harvested at each time point and directly lysed with sodium dodecyl sulfate-polyacrylamide gel electrophoresis loading dye for Western blot analysis using anti-phospho-ERK2 (Cell Signaling Inc.) or anti-pan-Ras antibodies (Oncogene Research Products). cAMP assay. cAMP levels were quantified by using a cAMP assay kit (Amersham). A total of 100 million cells were pulsed with 50 nM cAMP for 5 h (20 × 106 cells/ml), washed, and resuspended with DB buffer at a density of 5 × 107 cells/ml. Before the activation, cells were preincubated with DB buffer containing 10 mM dithiothreitol for 10 min on ice. A 1.8-ml aliquot of cells was transferred into a cup on an orbital shaker for 2 min. Then, 200 μl of stimulation cocktail (10 mM dithiothreitol and 100 μM of 2-deoxy cAMP) was added to the cells. Samples of 100 μl were taken at each time point into 1.5-ml Eppendorf tubes filled with 100 μl of 3.5% perchloric acid. Samples were mixed thoroughly and incubated on ice for 30 min. A 100-μl volume of 50% saturated KHCO3 solution was added to the sample and incubated on ice for 30 min. After insoluble aggregates were removed by centrifugation, the supernatants were rapidly frozen using liquid nitrogen. Samples were evaporated in a Speed-Vac without heating for 15 h and resuspended with 110 μl of Tris-EDTA buffer from the cAMP kit. Levels of cAMP were determined by following the manufacturer's instructions (Amersham). MPL1 is an active phosphatase with leucine-rich repeats. A group of genes encoding a potential DSPase domain with LRRs can be found in the genome of Dictyostelium and several protozoans, such as Entamoeba histolytica, Leishmania major, and Trypanosoma cruzi (Fig. ). These hypothetical genes show a potential DSPase (PPase) and LRR domains and sometimes extra domains, such as tandem serine/threonine kinase domains and/or Zn finger domains. No other similarly structured genes were found in the currently available genomes of metazoa or plants. We have cloned one such Dictyostelium gene, MPL1, which contains a carboxyl-terminal phosphatase domain with conserved residues essential for dual specificity phosphatase function and six potential leucine-rich repeats at the amino-terminal half of the protein (Fig. ). The MPL1 expression pattern was determined by Northern blot analysis using a probe encoding the phosphatase domain, as shown in Fig. . MPL1 expression was significantly up-regulated upon starvation, reaching a maximum around 10 h. MPL1 expression subsequently declined but was present throughout development. To ensure that MPL1 is an active phosphatase, we generated a GST fusion of the putative MPL1 phosphatase domain (MPL1-PPase; amino acids 580 to 834). GST and GST-MPL1-PPase were expressed and purified from Escherichia coli, using glutathione-Sepharose beads. Equivalent molar amounts of GST and GST-MPL1-PPase were normalized by Western blotting using anti-GST antibody (Fig. ), and equimolar amounts of the protein were incubated with pNPP. GST-MPL1-PPase clearly demonstrated a phosphatase activity of pNPP dephosphorylation (Fig. ). It is, however, not known if MPL1 is a bona fide dual-specificity phosphatase. mpl1− cells are defective in aggregation. mpl1− cells were created by homologous recombination as shown in Fig. . Ablation of the MPL1 gene was screened by PCR analysis and genomic Southern blotting using an MPL1-specific probe. To determine the level of a partial MPL1 transcript in mpl1− cells, transcripts were analyzed by RT-PCR using a primer set upstream of the blasticidin cassette insertion point. Compared to wild-type cells, mpl1− cells exhibited a significant decrease in the level of the partial MPL1 transcript, while Ig7 control transcript levels were comparable (Fig. ). Although faintly visible, the residual level of the partial MPL1 transcript did not interfere with the rescue of mpl1− phenotypes by the reintroduction of the full-length MPL1 gene into mpl1− cells (see Fig. to and Tables 1 to 3). The aggregation minus (agg-minus) phenotype was evident from mpl1− cells developed on Klebsiella aerogenes plates for seven days (Fig. ). Wild-type cells formed a number of fruiting bodies, whereas most of the mpl1− cells failed to display visible structures on the plate. Occasionally, in less than 10% of plaques, mpl1− cells formed various heterogeneous structures, including aggregates and fruiting bodies. This agg-minus phenotype was rescued by the reintroduction of the full-length MPL1 (Fig. ). | TABLE 1.Chemotactic indices with 0.1 μM cAMPa |
| TABLE 3.Movement of cells in the absence of cAMPa |
To determine whether mpl1− cells are defective in cell migration, cells were plated to form territorial streams at various cell densities, as described for Fig. . While wild-type cells displayed eminent territorial streaming, even at a density of 1.25 × 104 cell/cm2, mpl1− cells, in contrast, consistently failed to form territorial streams, even at a 20 times higher cell density. mpl1− cells occasionally formed loose aggregates, which might have been formed by nearby cells coalescing together in the absence of directional cell migration. mpl1− cells are defective in chemotaxis. mpl1− cells were challenged with weak (0.1 μM) and strong (2 μM) gradients of cAMP for a duration of 60 min to fully examine their behavior. Cells were made responsive to the cAMP gradient by 4 h of cAMP pulses and were challenged for 1 hour with a cAMP gradient formed from a micropipette filled with 0.1 μM or 2 μM cAMP. During the initial 20 min, mpl1− cells exhibited severely compromised gradient sensing and reduced speed (Fig. and ; Tables 1 and 2) compared with wild-type cells (Fig. and and Tables 1 and 2). During the last 20 min, the chemotaxis index of mpl1− cells improved ~75% compared to the wild type under either condition, but the degrees of improvement in motility were less (Tables 1 and 2). Although statistically meaningful ( P= 0.012), the difference of motility under weak and strong gradients was modest. Behaviors of mpl1− cells reintroduced with the full-length MPL1 were determined in an equivalent manner. Under 0.1 μM cAMP, these cells displayed a wild-type-like chemotaxis index and motility (Fig. and Table 1). Behaviors of HS174 cells ( 27), which lack ERK2, were monitored for comparison with other cell types described in this report. HS174 cells displayed more severe defects in gradient sensing and directional motility than mpl1− cells under both gradients (Tables 1 and 2). These results were obtained from three independent experiments, and the average values are summarized in Tables 1 and 2. | TABLE 2.Chemotactic indices with 2.0 μM cAMPa |
To determine if MPL1 function is necessary for general motility, cells were stimulated with cAMP pulses in the chemotaxis assay, and their movement was monitored in the absence of the cAMP gradient. Wild-type cells, mpl1− cells, mpl1− cells reintroduced with the full-length MPL1, and HS174 cells were pulsed as described earlier and were analyzed by tracing their movement for 30 min at 1-min intervals (Fig. ; Table 3). Compared to wild-type cells, mpl1− cells consistently displayed compromised random motility (~60%), whereas mpl1− cells reintroduced with the full-length MPL1 showed no significant difference from wild-type cells (Table 3). Cells lacking ERK2 seemed to have more problems than cells suffering from aberrant ERK2 phosphorylation (Table 3; Fig. ). These results showed that mpl1− cells were compromised in both random and directional motility, and reintroduction of MPL1 significantly restored both defects. MPL1 regulates poststimulus ERK2 adaptation. Considering that one of the major dual specific phosphatases are MKPs, we reasoned that mpl1− cells may experience aberrant ERK2 phosphorylation. ERK2 phosphorylation in response to cAMP stimulation was tested by Western blot analysis using an anti-phospho-ERK2-specific antibody (Cell Signaling Inc.) ( 1). Cells were stimulated with pulsatile cAMP for 4 h, treated with 2 mM caffeine for 30 min, and then challenged with 0.1 μM of cAMP. Under these conditions, in wild-type cells ERK2 phosphorylation was virtually undetectable before cAMP stimulation and reached its maximum at 1 min after the stimulation. mpl1− cells, however, not only displayed higher prestimulus ERK2 phosphorylation but also persistent phosphorylation after cAMP stimulation over 4 min (Fig. ). The same blot was stripped and reprobed with anti-pan-Ras antibody to confirm equal loading across the lanes. These data indicate that MPL1 regulates both prestimulus and poststimulus ERK2 phosphorylation. Consistently, reintroduction of full-length MPL1 lowered prestimulus ERK2 phosphorylation to an undetectable level, and the poststimulus level of phospho-ERK2 was similarly restored to that of the wild type (Fig. ). Western blotting using anti-flag antibody showed the expression of the full-length MPL1 (Fig. ). One of the well-established phenotypes of erk2− cells is their inefficiency in cAMP generation in response to cAMP receptor activation. This is, however, not due to the defective activation of adenylyl cyclase, but rather due to the excessive activity of RegA, a cytoplasmic phosphodiesterase. Upon receiving cAMP stimulation, wild-type cells typically display the initial 2 minutes of the slower phase of cAMP production, which is missing in regA− cells ( 19). This 2-minute window coincides with the transient ERK2 activation. It is, therefore, plausible that mpl1− cells also display lack of the initial 2-minute slower phase of cAMP production in response to receptor stimulation. Production of cAMP from wild-type and mpl1− cells was analyzed in a [H 3]cAMP competition assay after cAMP pulsing. The typical 2-minute slower phase in cAMP production was lost in mpl1− cells (Fig. ), similar to that of the regA− cells ( 19). Furthermore, reintroduction of the full-length MPL1 restored the cAMP production pattern comparable to that of the wild type (Fig. ), indicating that MPL1 is also essential for proper cAMP production. Phosphatases with architectures similar to MPL1 exist in other unicellular eukaryotes. Although multiple MKPs and DSPases exist in mammals, none of the mammalian dual specificity phosphatases contains LRR domains. The MKPs interact with MAP kinases either directly through the phosphatase domain or indirectly through an MKB domain. No MKB domain was found in the Dictyostelium genome by NCBI homology domain search. There exist, however, several potential DSPases equipped with LRR domains in the genome of certain unicellular eukaryotic organisms, such as Entamoeba histolytica, Leishmania major, and Trypanosoma cruzi as described in Fig. . All three of these organisms are potentially pathogenic, and thus understanding MPL1 function may shed new insight into the biology of these parasitic organisms. It will be interesting to determine if the MPL1-like genes of Entamoeba histolytica, which are highly homologous to MPL1 in domain organization, are also essential for ERK2 regulation and motility. MPL1 is an essential regulator of ERK2 function. In this report, we showed that MPL1 is essential for proper ERK2 regulation. Considering that multiple phosphatases exist in vivo, it is significant to observe aberrantly high levels of phospho-ERK2 in mpl1− cells. In addition, the kinetics of cAMP production of mpl1− cells resembled that of the regA− cells, further supporting the notion that mpl1− cells have higher ERK2 activity (Fig. ). Furthermore, reintroduction of MPL1 in mpl1− cells restored ERK2 phosphorylation and cAMP production similar to that of the wild type (Fig. ). It can be said that MPL1 is essential for proper ERK2 regulation, although it is not clear whether MPL1 directly or indirectly dephosphorylates phospho-ERK2. In addition to RegA regulation, ERK2 has also been implicated in polarized cytoskeletal reorganization ( 27). Upon receiving cAMP stimulation during the aggregation process ( 24), cells lose polarity by disintegrating the leading edge, become rounded up within a minute, and regain polarity by reforming a leading edge after 6 to 7 min. Cells lacking ERK2 fail to form the dominant F-actin-filled leading edge. Instead, erk2− cells generate crown-like structures filled with both F-actin and myosin II. It seems, therefore, that transient activation of ERK2 during the early response period is essential for selective assembly of F-actin from myosin II in the leading edge after 6 min of signal integration. Consistently, an elevation of prestimulus ERK2 phosphorylation and persistent poststimulus ERK2 phosphorylation in mpl1− cells were paralleled with defective motility. Furthermore, the restoration of normal ERK2 regulation in mpl1− cells by reintroduction of MPL1 significantly improved random motility and directional motility. It seems conceivable that not only the deprivation, but also excessiveness, of ERK2 activity could hamper dynamic F-actin and myosin II remodeling during cell movement. However, due to the short list of ERK2 substrates in Dictyostelium cells, it remains to be determined how deprivation of or excess ERK2 activity regulates cytoskeletal remodeling during cell movement. mpl1− cells were defective in cell migration. Quantitative analysis of the chemotaxis of mpl1− cells revealed several defects. Compared to the wild type, mpl1− cells displayed severely compromised chemotaxis index during the initial 20 min, which improved significantly when allowed for an additional 40 min under both weak and strong cAMP gradients. Considering their modest phenotype and the delayed improvement in directionality of mpl1− cells, it is unlikely that MPL1 is the major determinant of gradient sensing. In contrast, motility was more severely compromised during the first 20 min under both gradients. During the last 20 min, the motility improved modestly under both gradients. The degree of improvement was slightly better under the weak than the strong cAMP gradient. mpl1− cells may experience more problems in ERK2 adaptation under a strong cAMP gradient, but the mechanism behind these observations is currently not clear. In any case, mpl1− cells displayed a more severe defect in motility than the directionality. Reintroduction of the full-length MPL1 in mpl1− cells significantly restored multiple defects in aggregation, chemotaxis, random motility, ERK2 regulation, and cAMP production, which underscored the significance of MPL1 in these processes. Marbelys Rodriguez is a RISE student supported by NIH. Sudhakar Veeranki is an FIU-Presidential fellow. We thank Lisa Schneper and Lorraine Acevedo for their technical help and Lisa Schneper for critical reading and discussions. 1. Brzostowski, J. A., and A. R. Kimmel. 2006. Nonadaptive regulation of ERK2 in Dictyostelium: implications for mechanisms of cAMP relay. Mol. Biol. Cell 174220-4227. [PubMed] 2. Ducruet, A. P., A. Vogt, P. Wipf, and J. S. Lazo. 2005. Dual specificity protein phosphatases: therapeutic targets for cancer and Alzheimer's disease. Annu. Rev. Pharmacol. Toxicol. 45725-750. [PubMed] 3. Dynes, J. L., A. M. Clark, G. Shaulsky, A. Kuspa, W. F. Loomis, and R. A. Firtel. 1994. LagC is required for cell-cell interactions that are essential for cell-type differentiation in Dictyostelium. Genes Dev. 8948-958. [PubMed] 4. Early, A., M. Gamper, J. Moniakis, E. Kim, T. Hunter, J. G. Williams, and R. A. Firtel. 2001. Protein tyrosine phosphatase PTP1 negatively regulates Dictyostelium STATa and is required for proper cell-type proportioning. Dev. Biol. 232233-245. [PubMed] 5. Enkhbayar, P., M. Kamiya, M. Osaki, T. Matsumoto, and N. Matsushima. 2004. Structural principles of leucine-rich repeat (LRR) proteins. Proteins 54394-403. [PubMed] 6. Gamper, M., P. K. Howard, T. Hunter, and R. A. Firtel. 1996. Multiple roles of the novel protein tyrosine phosphatase PTP3 during Dictyostelium growth and development. Mol. Cell. Biol. 162431-2444. [PubMed] 7. Gamper, M., E. Kim, P. K. Howard, H. Ma, T. Hunter, and R. A. Firtel. 1999. Regulation of Dictyostelium protein-tyrosine phosphatase-3 (PTP3) through osmotic shock and stress stimulation and identification of pp130 as a PTP3 substrate. J. Biol. Chem. 27412129-12138. [PubMed] 8. Howard, P. K., B. M. Sefton, and R. A. Firtel. 1992. Analysis of a spatially regulated phosphotyrosine phosphatase identifies tyrosine phosphorylation as a key regulatory pathway in Dictyostelium. Cell 71637-647. [PubMed] 9. Howard, P. K., M. Gamper, T. Hunter, and R. A. Firtel. 1994. Regulation by protein-tyrosine-phosphatase Ptp2 is distinct from that by Ptp1 during Dictyostelium growth and development. Mol. Cell. Biol. 145154-5164. [PubMed] 10. Huang, Y. Z., M. W. Zang, W. C. Xiong, Z. J. Luo, and L. Mei. 2003. Erbin suppresses the MAP kinase pathway. J. Biol. Chem. 2781108-1114. [PubMed] 11. Keyse, S. M. 2000. Protein phosphatases and the regulation of mitogen-activated protein kinase signalling. Curr. Opin. Cell Biol. 12186-192. [PubMed] 12. Kobe, B., and A. V. Kajava. 2001. The leucine-rich repeat as a protein recognition motif. Curr. Opin. Struct. Biol. 11725-732. [PubMed] 13. Kolch, W. 2000. Meaningful relationships: the regulation of the Ras/Raf/MEK/ERK pathway by protein interactions. Biochem. J. 351289-305. [PubMed] 14. Kosaka, C., and C. J. Pears. 1997. Chemoattractants induce tyrosine phosphorylation of ERK2 in Dictyostelium discoideum by diverse signaling pathways. Biochem. J. 324347-352. [PubMed] 15. Laub, M. T., and W. F. Loomis. 1998. A molecular network that produces spontaneous oscillations in excitable cells of Dictyostelium. Mol. Biol. Cell 93521-3532. [PubMed] 16. Loftus, B., I. Anderson, R. Davies, U. C. Alsmark, J. Samuelson, P. Amedeo, P. Roncaglia, M. Berriman, R. P. Hirt, B. J. Mann, T. Nozaki, B. Suh, M. Pop, M. Duchene, J. Ackers, E. Tannich, M. Leippe, M. Hofer, I. Bruchhaus, U. Willhoeft, A. Bhattacharya, T. Chillingworth, C. Churcher, Z. Hance, B. Harris, D. Harris, K. Jagels, S. Moule, K. Mungall, D. Ormond, R. Squares, S. Whitehead, M. A. Quail, E. Rabbinowitsch, H. Norbertczak, C. Price, Z. Wang, N. Guillén, C. Gilchrist, S. E. Stroup, S. Bhattacharya, A. Lohia, P. G. Foster, T. Sicheritz-Ponten, C. Weber, U. Singh, C. Mukherjee, N. M. El-Sayed, W. A. Petri, Jr., C. G. Clark, T. M. Embley, B. Barrell, C. M. Fraser, and N. Hall. 2005. The genome of the protist parasite Entamoeba histolytica. Nature 433865-868. [PubMed] 17. Loovers, H. M., M. Postma, I. Keizer-Gunnink, Y. E. Huang, P. N. Devreotes, and J. M. van Haastert Peter. 2006. Distinct roles of PI(3,4,5)P3 during chemoattractant signaling in Dictyostelium: a quantitative in vivo analysis by inhibition of PI3-kinase. Mol. Biol. Cell 171503-1513. [PubMed] 18. Lyon, M. A., A. P. Ducruet, P. Wipf, and J. S. Lazo. 2002. Dual-specificity phosphatases as targets for antineoplastic agents. Nat. Rev. Drug Disc. 1961-976. 19. Maeda, M., S. Lu, G. Shaulsky, Y. Miyazaki, H. Kuwayama, Y. Tanaka, A. Kuspa, and W. F. Loomis. 2004. Periodic signaling controlled by an oscillatory circuit that includes protein kinases ERK2 and PKA. Science 304875-878. [PubMed] 20. Rintelen, F., E. Hafen, and K. Nairz. 2003. The Drosophila dual-specificity ERK phosphatase DMKP3 cooperates with the ERK tyrosine phosphatase PTP-ER. Development 1303479-3490. [PubMed] 21. Roux, P. P., and J. Blenis. 2004. ERK and p38 MAPK-activated protein kinases: a family of protein kinases with diverse biological functions. Microbiol. Mol. Biol. Rev. 68320-344. [PubMed] 22. Sawai, S., P. A. Thomason, and E. C. Cox. 2005. An autoregulatory circuit for long-range self-organization in Dictyostelium cell populations. Nature 433323-326. [PubMed] 23. Theodosiou, A., and A. Ashworth. 2002. MAP kinase phosphatases. Genome Biol. 3REVIEWS3009.1-3009.10. doi: doi: 10.1186/gb-21002-3-7-reviews3009.. [PubMed] 24. Tomchik, K. J., and P. N. Devreotes. 1981. Adenosine 3′,5′-monophosphate waves in Dictyostelium discoideum: a demonstration by isotope dilution-fluorography. Science 212443-446. [PubMed] 25. Tsai, H. R., L. M. Yang, W. J. Tsai, and W. F. Chiou. 2004. Andrographolide acts through inhibition of ERK1/2 and Akt phosphorylation to suppress chemotactic migration. Eur. J. Pharmacol. 49845-52. [PubMed] 26. Vindis, C., D. P. Cerretti, T. O. Daniel, and U. Huynh-Do. 2003. EphB1 recruits c-Src and p52 (Shc) to activate MAPK/ERK and promote chemotaxis. J. Cell Biol. 162661-671. [PubMed] 27. Wang, Y., J. Liu, and J. E. Segall. 1998. MAP kinase function in amoeboid chemotaxis. J. Cell Sci. 111373-383. [PubMed]
|
