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Originally published In Press as doi:10.1074/jbc.M609990200 on December 8, 2006

J. Biol. Chem., Vol. 282, Issue 7, 4884-4893, February 16, 2007
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ROCK1 Phosphorylates and Activates Zipper-interacting Protein Kinase*Formula

Laura Hagerty{ddagger}1, Douglas H. Weitzel{ddagger}1, Jenica Chambers{ddagger}, Christopher N. Fortner{ddagger}, Matthew H. Brush{ddagger}, David Loiselle{ddagger}, Hiroshi Hosoya§, and Timothy A. J. Haystead{ddagger}2

From the {ddagger}Department of Pharmacology and Cancer Biology, Duke University Medical Center, Durham, North Carolina 27710 and §Department of Biological Sciences, Graduate School of Science, Hiroshima University, Hiroshima 739 8526, Japan

Received for publication, October 24, 2006 , and in revised form, December 6, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Zipper-interacting protein kinase (ZIPK) regulates Ca2+-independent phosphorylation of both smooth muscle (to regulate contraction) and non-muscle myosin (to regulate non-apoptotic cell death) through either phosphorylation and inhibition of myosin phosphatase, the myosin phosphatase inhibitor CPI17, or direct phosphorylation of myosin light chain. ZIPK is regulated by multisite phosphorylation. Phosphorylation at least three sites Thr-180, Thr-225, and Thr-265 has been shown to be essential for full activity, whereas phosphorylation at Thr-299 regulates its intracellular localization. Herein we utilized an unbiased proteomics screen of smooth muscle extracts with synthetic peptides derived from the sequence of the regulatory phosphorylation sites of the enzyme to identify the protein kinases that might regulate ZIPK activity in vivo. Discrete kinase activities toward Thr-265 and Thr-299 were defined and identified by mass spectrometry as Rho kinase 1 (ROCK1). In vitro, ROCK1 showed a high degree of substrate specificity toward native ZIPK, both stoichiometrically phosphorylating the enzyme at Thr-265 and Thr-299 as well as bringing about activation. In HeLa cells, coexpression of ZIPK with ROCK1 altered the ROCK-induced phenotype of focused stress fiber pattern to a Rho-like phenotype of parallel stress fiber pattern. This effect was also dependent upon phosphorylation at Thr-265. Our findings provide a new regulatory pathway in smooth muscle and non-muscle cells whereby ROCK1 phosphorylates and regulates ZIP kinase.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The major mechanism governing the activity of myosin in smooth muscle and in many non-muscle cells is phosphorylation of its 20-kDa regulatory light chain (LC20).3 In vivo the level of LC20 phosphorylation is dependent upon the opposing activities of the Ca2+/calmodulin-dependent protein kinase, myosin light chain kinase, and myosin light chain phosphatase (SMPP-1M) (for reviews, see Refs. 1-3). Smooth muscles put considerable effort into regulating the respective activities of myosin light chain kinase and SMPP-1M. Inhibiting or activating either enzyme results in a net increase or decrease in LC20 phosphorylation and hence changes the contractile state of smooth muscle. We and others have hypothesized that regulation of SMPP-1M activity enables exquisite control of smooth muscle contractile state independently of [Ca2+] (1-4). In nonmuscle cells, regulation of myosin phosphorylation has been implicated in cytoskeletal reorganization, trafficking, and non-apoptotic cell death (3).

In smooth muscle SMPP-1M is regulated through phosphorylation of its myosin-targeting subunit MYPT1 at Ser-695/Thr-696 (numbering is based on the mammalian isoforms) and Thr-853 (for reviews, see Refs. 4 and 5). Phosphorylation at Thr-696 and Thr-853 inactivate the phosphatase toward myosin, sensitizing the muscle to the contractile effects of Ca2+ (Ca2+ sensitization) (6-13). Phosphorylation at Ser-695 has no direct effect on enzymatic activity but prevents phosphorylation at Thr-696 (14-16).

Several lines of evidence place Rho kinase (ROCK) as a primary regulator of MYPT1 in vivo (for a review, see Ref. 3). Addition of GTP{gamma}S to permeabilized smooth muscles induces profound Ca2+ sensitization, mimicking the actions of many agonists that exert their effects on smooth muscles through G protein-coupled receptors (17-21). RhoA is the major GTP-binding protein in smooth muscle (20), and subsequently it has been established that the small G protein plays an essential role in the process of Ca2+ sensitization in smooth muscles by inhibiting SMPP-1M activity. A major target for RhoA in smooth muscle is ROCK (22, 23). In vitro ROCK phosphorylates MYPT1 at both Thr-853 and Thr-696, causing dissociation of the phosphatase complex from myosin as well as inactivating the enzyme, but shows a preference for Thr-853 relative to Thr-696 (6-9, 11, 24-26). Treatment of isolated smooth muscles with the ROCK inhibitor Y-27632 selectively blocks contraction in response to Ca2+-sensitizing agonists (24, 25). Furthermore Y-27632 has been shown to dramatically correct hypertension in hypertensive rat models (25). Other lines of evidence may cast doubt on a direct involvement for ROCK in regulation of MYPT1. First, the recruitment of ROCK to RhoA-GTP at the cell membrane raises both temporal and spatial concerns about the access of the kinase to myosin-bound MYPT1 (27, 28). Second, we and other groups have identified several additional MYPT1 kinases that phosphorylate MYPT1 at Thr-696/853 in smooth muscle and in other cell types. These kinases include ZIPK, identified by our group (29); integrin-linked kinase (29); and myotonic dystrophy kinase (29). Third and perhaps most intriguingly, neither the ROCK1 nor the ROCK2 null mouse has an obvious phenotype that would suggest a role in the regulation of smooth muscle contractility (30, 31).

Our laboratory has focused on ZIPK as a primary regulator of MYPT1 and myosin phosphorylation in smooth muscle. Evidence supporting a role for ZIPK in the regulation of smooth muscle contraction includes the following. 1) ZIPK co-purifies with MYPT1 and myosin in smooth muscle (29). 2) ZIPK was identified as the primary binding target for MYPT1 in an unbiased high stringency two-hybrid screen using MYPT1 as bait (32). 3) ZIPK is the major MYPT1 kinase activity present in muscle extracts (29). 4) ZIPK phosphorylates MYPT1 at Thr-696 and Thr-853 in vitro. 5) MYPT1 exhibits a low Km (~2 µM) for ZIPK (29). 6) Addition of recombinant ZIPK to permeabilized smooth muscle causes profound Ca2+ sensitization (29, 33). 7) ZIPK phosphorylates the SMPP-1M inhibitor CPI17 as well as myosin light chain in vitro (29, 33).

ZIPK is itself regulated by phosphorylation. We recently identified six phosphorylation sites on the protein: three that are required for enzymatic activity (Thr-180, Thr-225, and Thr-265), one that regulates nuclear localization (Thr-299), and two others of unknown function (Thr-306 and Thr-311) (34). Prior to our work, Kimchi and co-workers identified six other potential ZIPK regulatory sites (Ser-306, Ser-309, Ser-311, Ser-312, Ser-318, and Ser-326) (35, 36). More recently, Sato et al. (37) demonstrated that ZIPK is phosphorylated at Thr-265 in non-muscle cells treated with interleukin 6. Prior to these studies, our group had shown that ZIPK is activated and phosphorylated in smooth muscle following treatment with the Ca2+-sensitizing agonist carbachol (29).

The nature of the protein kinases that regulate ZIPK activity in smooth muscle are not known. Overexpression of at least one kinase in HEK293 cells, DAPK1, has been reported to activate ZIPK (36). Herein we used an unbiased proteomics screen to characterize major ZIP kinase kinase activities in smooth muscle and identified ROCK1 as a major Thr-265/Thr-299 peptide kinase. In vitro ROCK1 stoichiometrically phosphorylates ZIPK at Thr-265 and Thr-299, resulting in an increase in ZIPK enzyme activity. In a series of transfection experiments, ZIPK altered the ability of constitutively active ROCK to promote cytoskeletal reorganization.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
The following peptide substrates were synthesized and purified by Dr. John Gorka (Biomolecules Midwest Inc., St. Louis, IL): peptide T180, KKKKNIFGTPEFVAPE; peptide T225, KKKQETLTNISAVNYD; peptide T265, KRRMTIAQSLEH-SWIK; peptide T299, RRRLKTTRLKEYTIK; peptide T306, KKKEYTIKSH; and peptide S311, KKSHSSLPPNNSYAD. Phosphospecific antibodies were prepared from phosphosynthetic peptides derived from the Thr-180, Thr-225, Thr-265, Thr-299, Thr-306, and Ser-311 phosphorylation sites of ZIPK by Proteintech Inc. All other antibodies were purchased from Calbiochem. Vectors expressing various forms of ZIPK were prepared as described previously (34). Recombinant full-length and constitutively active forms of ROCK1 were either purchased from Signal Transduction Laboratories or purified as FLAG fusion proteins from transfected HEK293 cells.

Analysis of ZIP Kinase Kinase Activity in Smooth Muscle—Whole porcine pig bladders (100 g) freshly isolated from male pigs were washed and incubated in Krebs buffer gassed with 95% O2, 5%CO2 at 37 °C and then treated for 5 min with 10 µM carbachol. The fully contracted bladders were removed from the medium, cut and stripped of their mucosa with a glass rod, and then plunged into a bath of liquid nitrogen. The frozen bladders were crushed into a powder under liquid N2 and homogenized in 2.5 volumes (w/v) of buffer A (4 mM EDTA, pH 7.2 at 25 °C, 1 mM dithiothreitol, 6 µg/ml leupeptin, 1 mM aprotinin, 1 mM benzamidine, 1 mM pepstatin, and 10 µM microcystin). The homogenate was centrifuged for 30 min at 30,000 x g, and the pellet and supernatant (cytosolic fraction) were separated. The pellet (myofibrillar fraction) was re-extracted with 500 ml of buffer B (50 mM Tris-HCl, pH 7.2 at 25 °C, 600 mM KCl, 0.2% Nonidet P-40, 1 mM dithiothreitol, and the mixture of protease and phosphatase inhibitors used in buffer A). After gentle stirring for 30 min at 4 °C, the myofibrillar fraction was centrifuged at 30,000 x g for 30 min. The cytosolic and myofibrillar extracts were mixed gently with 10 ml of {gamma}-phosphate-linked ATP medium in the presence of 60 mM MgCl2 for 30 min (38). The medium was washed with buffer B containing 1 M KCl and 60 mM MgCl2 and then incubated for 30 min with 20 ml of buffer C (buffer B containing 100 mM ATP, 150 mM KCl, and 60 mM MgCl2). The column eluate was collected, rapidly concentrated 3-fold to 1 ml by a Centricon filter (3000 x g using a 10,000-Da cutoff filter for 20 min each) to reduce the ATP concentration to <100 µM. The concentrated eluate was applied to a micro 0.5 x 1.5-cm Mono Q anion-exchange column equilibrated in buffer D (25 mM Tris-HCl, pH 7.2 at 25 °C, 1 mM dithiothreitol). The bound proteins were eluted over 90 min with a salt gradient to 1 M NaCl in buffer D. Fractions (50 µl) were diluted 10-fold with buffer D and assayed for protein kinase activity. Greater than 95% of the total Thr-180, Thr-265, and Thr-299 peptide kinase activity measured was found in the myofibrillar fraction. Therefore, all subsequent studies were carried out on this fraction. Active column fractions from the myofibrillar fraction were analyzed by SDS-PAGE and silver staining. Silver-staining proteins corresponding to peptide kinase activity were excised from the gel and treated with trypsin, and proteins were identified in an Applied Biosystems 4700 matrix-assisted laser desorption ionization time-of-flight time-of-flight (MALDI-TOF-TOF) mass spectrometer as described previously (34).

Protein Kinase Assays—Reactions for protein kinase activity contained 10 µl of diluted fraction and 10 µl of peptide substrate (prepared from 2.5 mg/ml solution in buffer B) and were initiated with 10 µl of 300 µM ATP, 2.5 mM MgCl2 (2500 cpm/nmol). After 5 min at 25 °C, the reactions were stopped with 20 µl of 200 mM phosphoric acid, and 10 µl of the reaction mixture were spotted on 1-cm2 segments of p81 paper (Whatman), washed in 200 mM phosphoric acid, and Cerenkov counted.


Figure 1
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  		FIGURE 1.
ZIP kinase kinase activities in smooth muscle extracts. Smooth muscle extracts were enriched for protein kinase activity by {gamma}-phosphate ATP affinity chromatography and then fractionated by microanion-exchange chromatography. In A, chromatography column fractions were assayed for protein kinase activity with the indicated peptides derived from the regulatory phosphorylation sites of ZIPK and MYPT1 in the presence and absence of 10 µM Y-27632. B shows the respective activities of recombinant ZIPK and ROCK1 against a peptide derived from the sequence surrounding Thr-265 on ZIPK. C shows the respective activities of purified recombinant ZIPK and ROCK1 against peptides containing the regulatory phosphorylation sites on MYPT1.

 


Figure 2
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  		FIGURE 2.
Proteomics analysis of anion-exchange column fractions containing ZIP kinase kinase activities. Anion-exchange column fractions containing the major peaks of Thr-265 and Thr-299 activities were characterized by SDS-PAGE and silver staining. Individual protein bands in each fraction were excised, digested with trypsin, and identified by mass spectrometry. 1 and 2, SPTA2; 3, SPTA2/MYH11; 4, PPP6; 5, MYH11, HS90A; 6, MYH11; 7, 8, 9, and 10, ROCK1; 11, HS90A; 12, TERA; 13, GEPH, ROCK2, GOGA4; 14, HS90A, TERA, GEPH; 15, endoplasmin precursor (94-kDa glucose-regulated protein), HS90A; 16, HS90A; 17, HS90A; 18, HS90A, TERA, GEPH; 19, 20, and 21, HS90A; 22, EHD2; 23, TCPG, TCPA1, TCPA2, ROCK2; 24, TCPA1, TCPG; 25, HS90A, TPM4; 26, TCPA; 27, TCPQ, TCPH; 28 and 29, TCPQ, HS90A; 30, PCCB, IMDH2; 31, HS90A; 32, PCCB, IMDH2; 33, HS90A; 34, PI52A; 35, PI52A, GEPH; 36, HS90A; 37, ACTH, adenosylhomocysteinase (S-adenosyl-L-homocysteine hydrolase); 38, HS90A, ACTH; 39, HS90A; 40, HS90A, ACTH; 41, ACTH; 42, HS90A; 43, TPM1; 44, glyceraldehyde-3-phosphate dehydrogenase; 45, 46, and 47, HS90A; 48, TCPG; 49 and 50, ABCF1; 51, 14-3-3 protein {zeta}/{delta} (protein kinase C), 14-3-3 protein {gamma} (protein kinase C). See supplemental data for complete raw MS data. LC, liquid chromatography.

 
Phosphopeptide Mapping—All mapping studies with ROCK1 were carried out by incubating 200 ng of recombinant ROCK1 with 1 µg of ZIPK in the presence of 100 µM [{gamma}-32P]ATP, 2.5 mM MgCl2 (2500 cpm/nmol) for 20 min at 25 °C. The 32P-labeled protein was separated by SDS-PAGE, silver-stained, and detected by autoradiography. The stoichiometry of phosphorylation was determined by excising the labeled protein, Cerenkov counting, and dividing the amount of radioactivity incorporated by the specific activity of ATP in the reaction. The precise sites were mapped by reverse phase HPLC and CRP analysis following extraction of proteolytic fragments from the gel as described previously (29, 34).

Cell Culture, Transfection, Purification of Recombinant ZIPK, and Fluorescence Microscopy—HeLa cells were maintained in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% fetal bovine serum. Protocols for transfection and purification of FLAG-ZIPK and the mutants used in this study are described in Graves et al. (34). For microscopic analysis, cells were plated onto glass coverslips and transfected using FuGENE (Roche Applied Science). Twenty-four hours after transfection, cells were washed once in phosphate-buffered saline, fixed in 4% paraformaldehyde (Sigma), and lysed in 0.2% Triton. Nonspecific sites were blocked with 1% bovine serum albumin in phosphate-buffered saline followed by incubation with AlexaFluor 568 phalloidin (Molecular Probes) according to the manufacturer's instruction. Images were obtained with a Zeiss LSM 410 confocal microscope.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Analysis of ZIP Kinase Kinase Activity within the Smooth Muscle Purinome—To characterize and identify ZIP kinase kinase activities in smooth muscle, a series of synthetic peptide substrates were derived from the primary amino acid sequence surrounding the major regulatory phosphorylation sites on ZIPK, Thr-180, Thr-265, and Thr-299. Additionally because ZIPK can autophosphorylate, assays were also carried out on a synthetic peptide derived from the amino acid sequence of Thr-696 from MYPT1 (29). A combination of ATP affinity and anion-exchange chromatography was used to rapidly separate out various kinase components within smooth muscle and more readily define ZIP kinase kinase activity. Fig. 1A shows that assay of anion-exchange column fractions with both Thr-265- and Thr-299-containing peptides identified a major peak of kinase activity eluting between fractions 32 and 40. In contrast, assay with MYPT1T696 identified two peaks of kinase activity, the first co-eluting Thr-265/Thr-299 activity and the second between fractions 42 and 48. Previously we have shown that the major MYPT1T696 activity present in smooth muscle extracts corresponds to two forms of ZIPK, a 54-kDa form and a 32-kDa proteolytic fragment (29). Assay with ZIPKT180 peptide characterized a small peak of activity eluting between fractions 32 and 40.

To identify the types of protein kinases co-eluting across fractions 32-40, the fractions were separated by one-dimensional SDS-PAGE and characterized by silver staining (Fig. 2). Fig. 2 shows that fractions 34-39 contained several proteins of varying molecular weight and abundance. To characterize the compliment of proteins contained in each gel lane, each silver-staining protein band was excised and digested with trypsin as described previously (39). Following extraction from the gel slices, tryptic peptide mixtures were analyzed by high through-put mass spectrometry using in an Applied Biosystems MALDI-TOF-TOF mass spectrometer. Because individual stained gel bands were likely to contain several proteins of varying abundance, the extracted tryptic peptides were analyzed first by peptide mass fingerprinting followed by extensive MS/MS analysis of all peptides identified with a signal to noise ratio >5. The compiled raw peptide mass fingerprinting and MS/MS data were searched against the public databases to identify their protein content (supplemental data I). To ensure unambiguous identification resulting from peptide mass redundancy or other known mass spectrometric related artifacts, stringent search criteria were applied, requiring multiple peptide matches within a given peptide mass fingerprint for an identified protein plus a minimum of two separate MS/MS matches. Using this approach 95 peptide mass fingerprinting spectra and 736 MS/MS spectra were collected identifying over 40 different proteins co-eluting throughout fractions 34-39. Importantly our MS analysis showed that many of the excised gel bands contained multiple proteins. For example band 11 contained three distinct proteins (Gephyrin, ROCK2, and Golgin 1) all showing multiple distinct peptide mass and MS/MS matches (see supplemental data I). Comparison of the average MS signal intensity of digests containing mixtures demonstrated that most proteins could be unambiguously identified over a differential concentration range of ~5-10-fold. These data demonstrate that our MS-based approach can be used to identify multiple proteins of varying abundance co-eluting within a single silver-staining band excised from an SDS gel.

Inspection of the list of proteins identified in Fig. 2 shows that the majority were members of the purine-binding protein superfamily or proteins known to bind to a purine-utilizing enzyme. Of interest to this study were bands that contained protein kinases that co-eluted with Thr-265, Thr-299, and Thr-180 activity. Although the entire chromatogram contained multiple protein kinases and purine-binding proteins (data not shown), ROCK1, ROCK2, and ZIPK were the only protein kinases positively identified in fractions corresponding to Thr-265, Thr-299, and Thr-180 kinase activity (region 32-40). In vitro purified recombinant ROCK and ZIPK demonstrated activity against the Thr-265 and Thr-299 peptides (supplemental data, Fig. 1B, and Table 1). In contrast, neither kinase showed activity toward Thr-180 peptide, suggesting that the Thr-180 kinase is likely to be distinct from those governing phosphorylation at Thr-265/Thr-299. The identity of the Thr-180 kinase is currently being investigated. Western analysis with antibodies to ROCK1 and ZIPK confirmed mass spectrometric data identifying both kinases with blotting intensity correlating with kinase activity shown in Fig. 1A (data not shown). Additionally in the case of ROCK1, analysis of the numbers of MS ions assigned and MS/MS data collected for ROCK1 for each fraction showed that these values were highest in fractions 35-37, fractions in which Thr-265/Thr-299 was also at its highest.

To discriminate ROCK1 activity from ZIPK, the fractions were reassayed against the Thr-265/Thr-299 peptides in the presence and absence of the ROCK1 inhibitor Y-27632 (10 µM). Fig. 1A shows that >90% of the activity directed toward either Thr-265 or Thr-299 peptides was inhibited. In contrast, activity measured using either the Thr-180 peptide or MYPT1T696 was only marginally affected by Y-27632 (data not shown). These findings strongly suggest that the majority of the observed Thr-265/Thr-299 kinase activity was due to ROCKI rather than ZIPK. They also rule out the possibility that the fractions contained other activities that contributed to Thr-265/Thr-299 activity. The lack of sensitivity of endogenous ZIPK (MYPT1T696) activity toward 10 µM Y-27632 is also consistent with previous results from our laboratory showing that ZIPK is relatively insensitive to the inhibitor (IC50 ZIPK = 340 µM) (29). To further confirm that the Thr-265/Thr-299 activities observed in the fractionated extracts were likely to be attributable to native ROCK and not ZIPK, we determined the relative Km and kcat values for the Thr-265 and Thr-299 peptides using active recombinant forms of each kinase (Table 1 and Fig. 1B). ROCK1 demonstrated a significantly higher specificity constant for both peptides relative to ZIPK (~8.5-fold greater for Thr-265 and ~9.3-fold greater for Thr-299). These data suggest that native ROCK1 is indeed responsible for the kinase activity toward ZIPK Thr-265 and Thr-299. Finally ROCK1 and ZIPK activity toward the Thr-265 and Thr-299 peptides were compared with their activity toward MYPT1T696 and MYPT1T853 peptides (Fig. 1C). Table 1 and Fig. 1, B and C, show that recombinant ROCK1 demonstrates a surprising ~1.7-fold preference for ZIPKT265 over even the MYPT1T853 peptide. In contrast, ZIPK exhibited a strong preference for MYPT1T696 relative to the MYPT1T853 or ZIPKT265 peptides.


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  		TABLE 1
The specificity of purified ZIPK and ROCK1 for the major regulatory phosphorylation sites on ZIPK and MYPT1

 
ROCK1 Phosphorylates ZIPK at Thr-265 and Thr-299 in Vitro—To determine whether full-length ZIPK was a substrate for ROCK1 in vitro, the following recombinant forms of FLAG-ZIPK were expressed in HEK293 cells and immunoaffinity purified: wild-type, T180A, D161A, T225A, T299A, and T265A ZIPK. Because the wild-type form of ZIPK can autophosphorylate, initial studies examining ROCK1 phosphorylation were carried out using the D161A "kinase-dead" form of ZIPK (34). Aspartic acid 161 is a highly conserved amino acid found in the active site of all Ser/Thr protein kinases and is required for catalysis of ATP (38). Fig. 3A shows an autoradiogram showing incorporation of [32P]orthophosphate into D161A ZIPK (MS analysis confirmed that the lower band in the autoradiogram is ZIPK, and the upper fainter band is the truncated form of autophosphorylated ROCK). Measurement of the stoichiometry of phosphorylation (~1.43 mol/mol) suggested that one to two sites may be targeted. To identify the precise phosphorylation sites 32P-labeled D161A ZIPK was digested with the indicated proteases, and labeled phosphopeptides were characterized by reverse phase HPLC (Fig. 3, B-D). Fig. 3B shows that digestion with trypsin identifies three major radiolabeled peptides (labeled 1-3). Each peptide was isolated and subjected to CRP analysis in an Applied Biosystems automated gas phase sequencer (29). This analysis enables radiolabeled phosphoamino acids to be positionally placed within a tryptic peptide relative to Arg and Lys. This strategy was chosen over an MS-based approach because previous work had shown that the majority of ZIPK regulatory phosphorylation sites were not directly amenable to MS analysis (34). Most of the sites upon proteolytic cleavage produce either highly charged or undesirable peptide fragments for electrospray or MALDI-based approaches. Fig. 3C (bar graph inset) shows that radioactivity was identified at cycle 4 for tryptic peptide 1, cycle 3 for tryptic peptide 2, and cycle 3 and cycle 5 for tryptic peptide 3. This analysis narrowed the possible number of potential ROCK1 phosphorylation sites to Thr-003, Ser-51, Ser-52, Ser-57, Thr-112, Thr-225, Ser-288, Thr-306, Ser-311, Thr-312, Ser-371, Thr-265, Thr-299, Thr-300, Ser-429 for tryptic peptides 1, 2, and 3 and to residues Thr-180, Thr-277, and Ser-373 for the additional cycle 5 release measured with tryptic peptide 3. Based upon previous work and the analysis with synthetic peptides described above, these findings strongly suggested that Thr-265 and Thr-299 are primary sites of phosphorylation by ROCK1. In other studies, peptides derived from Thr-180, Thr-225, Thr-306, and Ser-311 were not recognized by purified ROCK1 in vitro making these sites unlikely candidates. To verify these conclusions, CRP analysis was repeated following cleavage of the phosphorylated protein with lysyl endopeptidase C (cleaves at Lys only). Phosphopeptide analysis by reverse phase HPLC identified two major peptides (Fig. 3D), and CRP analysis identified release of radioactivity at cycle 8 for Lys-peptide 1 and cycle 4 for Lys-peptide 2 (Fig. 3E). These findings suggest that tryptic peptide 3 and Lys-peptide 1 are derived from the Thr-299 site, and tryptic peptide 1/2 and Lys-peptide 2 are derived from Thr-265. This analysis also explains the chromatogram shown in Fig. 1 in which three major peptides are identified rather than two. Inspection of the Thr-265 site shows that it contains a series of Lys and Arg residues. The release of radioactivity at cycle 3 (tryptic 1) and cycle 4 (tryptic 2) is most likely explained by alternate cleavage of immediately adjacent Arg and Lys, 3 and 4 residues away from Thr-265. To confirm site analysis further, phosphorylation experiments were repeated using the T265A and T299A ZIPK mutants following phosphorylation in vitro with ROCK1 (Fig. 3D). In the case of the T265A mutant, only one peptide was identified (Lys-peptide 1) following selective cleavage at Lys. This peptide also released radio-activity at cycle 8 consistent with Thr-299. Phosphopeptide mapping of the T299A mutant is somewhat more complicated because additional peptides are present (Fig. 3D). In previous work, although the T299A mutation alters the cellular location of ZIPK, it does not inhibit its enzymatic activity or ability to autophosphorylate at Thr-180, Thr-265, Thr-225, Thr-306, and Ser-311 (34). However, the chromatogram does show absence of radioactivity at the Lys-peptide 1 position. It should also be noted that the Thr-299 site contains a Lys residue immediately N-terminal to Thr-299, explaining why Lys-peptide 1 yielded radioactivity at cycle 8 and not cycle 1. Typically when a phosphoamino acid occurs immediately adjacent to an Arg or Lys, this renders the site resistant to proteolytic cleavage.


Figure 3
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  		FIGURE 3.
Characterization of the primary ROCK1 phosphorylation sites on ZIPK. The D161A mutant form of full-length recombinant ZIPK was phosphorylated to ~1.4 mol/mol with purified ROCK1. A shows an auto-radiogram of the phosphorylated protein; recombinant ROCK1 was identified in the upper band, and recombinant ZIPK was identified in the lower band. In B the ZIPK band was digested with trypsin, and the 32P-labeled phosphopeptides were characterized by reverse phase HPLC and Cerenkov counting of column fractions. In C isolated phosphopeptides were characterized by CRP analysis as described under "Experimental Procedures." In D and E 32P-labeled ZIPK was digested with lysyl endopeptidase C and characterized by reverse phase HPLC and CRP analysis as described in Fig. 1, B and C.

 


Figure 4
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  		FIGURE 4.
ROCK1 phosphorylates and activates ZIPK in vitro. In A, a series of phosphospecific antibodies were derived to the indicated phosphorylation sites on ZIPK. The antibodies were tested for specificity against autophosphorylated ZIPK and recombinant ZIPK with the indicated Ser/Thr -> Ala substitutions. In B phosphorylation of Thr-265 and Thr-299 was measured in vitro following phosphorylation of D161A ZIPK with increasing concentrations of ROCK1. In C, the activity of wild type (WT), T225A, and T265A was measured against the MYPT1T696 peptide following preincubation with the indicated amounts of ROCK. The preincubation assay was halted after 10 min by addition of 10 µM Y-27632.

 
Final confirmation of Thr-265 and Thr-299 as the primary ROCK1 phosphorylation sites on ZIPK was obtained with phosphospecific antibodies developed to recognize these two sites (Fig. 4). Fig. 4A shows the specificity for four phosphospecific antibodies that selectively recognize Thr-225, Thr-265, Thr-299, and Ser-311 on recombinant wild-type ZIPK following autoactivation but not Thr/Ser -> Ala mutations of these respective sites. Fig. 4B shows that increasing concentrations of ROCK1 induce site-specific phosphorylation at Thr-265 and Thr-299 consistent with phosphopeptide mapping studies. Specificity of ROCK1 for Thr-265 and Thr-299 may be explained by the presence of a cluster of 3-4 basic amino acids +1or +2 residues N-terminal to the phosphorylated amino acid that is notably absent in all other ZIPK phosphorylations sites.

ROCK1 Activates ZIPK through Phosphorylation of Thr-265 in Vitro—Previous work by our laboratory established that ZIPK Thr-265 phosphorylation is critical for enzyme activity (34). Therein the ZIPK T265A mutant was shown to be inactive toward MYPT1T696 as well as LC20S19 peptide substrates. Additionally the Thr-265 mutant showed markedly diminished autophosphorylation. To determine whether ROCK1 phosphorylation of Thr-265 promotes ZIPK activity, we pretreated ZIPK and the T265A mutant with increasing concentrations of ROCK1. ZIPK activity toward the MYPT1T696 peptide substrate was isolated by inhibiting ROCK1 with 10 µM Y-27632 following the 10-min pretreatment. As shown in Fig. 4C, ZIPK activity increased dramatically following pretreatment with ROCK1. Furthermore the effect of ROCK1 on ZIPK activity was concentration-dependent. In contrast, the ZIPK T265A mutant remained inactive following pretreatment with up to 200 ng of ROCK1. These data establish that ROCK1 activates ZIPK and that this activation results from phosphorylation at Thr-265.

Previous studies by our laboratory determined that phosphorylation at Thr-225 was also required for ZIPK activity. The ZIPK T225A mutant, like the T265A mutant, showed minimal activity toward peptide substrates and minimal autophosphorylation (34). Intriguingly we have demonstrated in the present study that the T225A mutant can attain full activity toward the MYPT1T696 peptide substrate following preincubation with ROCK1 (Fig. 4C). The ZIPK T225A mutant becomes activated by ROCK1 in a concentration-dependent manner, although full activation of T225A requires higher concentrations of ROCK1 relative to wild-type ZIPK. This is presumably due to the inability of ZIPK T225A to autophosphorylate at Thr-265 and autoactivate. These data establish that phosphorylation at Thr-265 can promote ZIPK activation independently of phosphorylation at Thr-225. They also indicate that ROCK1 phosphorylation at Thr-265 is sufficient to produce full activity of ZIPK in vitro.

ROCK1 Signals via ZIPK in Intact Cells—We have previously shown that in vivo ZIPK activity from smooth muscle was inhibited by Y-27632. This finding contrasts with in vitro observations showing that ZIPK was insensitive to Y-27632, suggesting that in vivo ZIPK activation may be downstream of ROCK signaling (29). In the present study, we performed transfection studies in HeLa cells using phalloidin staining of F-actin structures as a marker of ROCK, Rho, and ZIPK activities. Activated forms of both Rho and ROCK have been shown to regulate actin stress fiber formation (40-42). Overexpression of RhoA 63L, a constitutively active mutant, produces a parallel array of stress fibers, whereas expression of activated ROCK{Delta}3, a truncated mutant lacking the Rho-binding domain, results in morphologically distinct, focused patterns of actin fibers (22, 40, 42, 43). Both wild-type and T265A ZIPK demonstrated little effect on the actin cytoskeleton when expressed alone (Fig. 5, A and B). However, coexpression of ROCK{Delta}3 and wild-type ZIPK resulted in a morphological alteration of stress fibers from a ROCK-like phenotype to a Rho-like phenotype (Fig. 5E). Coexpression with T265A ZIPK did not modulate the ROCK{Delta}3-induced focused actin fibers, under-scoring the importance of the Thr-265 activation site on ZIPK (Fig. 5F). Quantification of focused actin fiber arrangements in transfected HeLa cell populations shows that strong ROCK{Delta}3 effects on the cytoskeleton can be altered by ZIPK coexpression as described above (Fig. 6). ZIPK T265A mutant coexpression resulted in a rescue of ROCK{Delta}3-induced stress fibers as compared with wild-type ZIPK, again highlighting the role Thr-265 may have in transmitting ROCK signals. Equal amounts of wild-type or T265A FLAG-ZIPK were present in cells cotransfected with ROCK{Delta}3 (Fig. 6, inset). A previous report demonstrated that coexpression of the actin-binding protein mDia with ROCK{Delta}3 resulted in a similar switching from the ROCK pattern to a Rho-like pattern (44). These results suggest that ZIPK is likely to act downstream of ROCK to distribute its signaling evenly over the actin cytoskeleton rather than promoting a focused concentration of stress fibers in one or a few sites, and such ZIPK action is dependent upon phosphorylation of its Thr-265 residue.


Figure 5
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  		FIGURE 5.
ZIPK coexpression modulates ROCK1-induced stress fibers. HeLa cells were transfected with pEGFP and FLAG-ZIPK (A), FLAG-ZIPK T265A (B), GFP-RhoA 63L (C), Myc-ROCK{Delta}3(D), Myc-ROCK{Delta}3 and FLAG-ZIPK (E), and Myc-ROCK{Delta}3 and FLAG-ZIPK T265A (F). The cells were then stained with phalloidin to visualize the filamentous actin. GFP, green fluorescent protein.

 


Figure 6
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  		FIGURE 6.
Quantification of focused stress fiber patterns. HeLa cells transfected with pEGFP and the indicated constructs were examined for ROCK{Delta}3-induced focused actin structures by phalloidin staining. Approximately 100 green fluorescent protein-positive cells were counted and evaluated for focused stress fiber patterns, and a ratio of focused stress fiber patterns per green fluorescent protein-positive cells was generated. Counts were performed in triplicate, averaged, and graphed with standard error. t test results demonstrate significant differences between pairs tested with p values <0.01 for each. * indicates ROCK/ZIPK compared with ROCK/ZIPK T265A. ** indicates ROCK compared with ROCK/ZIPK T265A.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Data presented herein strongly suggest that ZIPK is a direct target of ROCK in vivo. In summary, we have shown the following. 1) Both native smooth muscle and recombinant ROCK selectively phosphorylate ZIPK at two of its regulatory sites (Thr-265 and Thr-299), both of which conform to ROCK consensus phosphorylation sites found in other known targets (including the activating 504/508 sites of LIM kinase (LIMKI/II) and inhibitory sites Thr-696 and Thr-853 of MYPT1). 2) ROCK activates both wild type and ZIPK T225A in vitro.3)In muscle extracts, ROCK accounts for >90% of the activity directed toward the Thr-265 and Thr-299 peptides. 4) ROCK{Delta}3-induced cytoskeletal reorganization is modified by ZIPK coexpression, and this modification is Thr-265 phosphorylation site-dependent.

Although ZIPK and ROCK clearly regulate myosin phosphorylation in both smooth muscle and non-muscle cells, surprisingly few laboratories have considered the possibility that these two enzymes may in fact directly interact (4, 29, 32). Several lines of evidence may already support this idea. First significant parallels exist between ROCK and ZIPK in both non-muscle and smooth muscle cells. In certain non-muscle cells (e.g. HEK293), when either kinase is overexpressed or expressed in constitutively activated states, both kinases cause cell rounding, blebbing, and cytoskeletal reorganization that is associated with myosin phosphorylation (34, 35, 43, 45, 46). In smooth muscle cells, both kinases cause Ca2+ sensitization. In smooth muscle ROCK phosphorylates many of the same substrates as ZIPK in vitro, including MYPT1, LC20, and CPI17 (11, 29, 33, 45, 47-49). Both kinases show a preference for threonine and target similar phosphorylation site consensus sequences, i.e. 3-4 basic amino acids +1or +2 N-terminal to the phosphorylation site. Previous work from our laboratory suggested that ROCK has an upstream role in the regulation of ZIPK activity. In smooth muscle, carbachol-induced activation of ZIPK was sensitive to Y-27632 even though ZIPK is not directly inhibited by this compound in vitro (29). More recently, Mendelsohn and co-workers (32) demonstrated that transfection of HEK293 cells with RhoA promoted association of ZIPK with MYPT1. However in smooth muscle, spatial and temporal issues suggested that ROCK may not directly mediate phosphorylation of these proteins in vivo. Native ROCK is largely membrane-bound when activated by RhoA, raising the question as to how ROCK can directly phosphorylate MYPT1 and myosin (27, 28). Although MYPT1 has been shown to localize to the membrane in smooth muscle cells (50, 51), this again begs the identity of the cytosolic kinase that induces MYPT1 phosphorylation and translocation. These previous findings in combination with data presented herein suggest that ZIPK stands squarely down-stream of both Rho and ROCK signaling, forming a signal transduction module to ultimately regulate myosin phosphorylation in both smooth muscle and non-muscle cells. A key question raised by studies herein, however, is the molecular mechanism by which ZIPK communicates with ROCK in vivo. Given the location of active ROCK at the plasma membrane, mechanisms must exist to translocate ZIPK to activated ROCK. One mechanism may be via binding to MYPT1. As discussed, others and we have shown that ZIPK binds directly to MYPT1 both in vitro and in vivo (4, 32). Studies by Hartshorne and co-workers (52) have demonstrated movement of MYPT1 from the cytoskeleton to the plasma membrane in intact cells. A possible mechanism, therefore, whereby ROCK directly interacts with ZIPK is via translocation of MYPT1 itself. Such a mechanism is attractive because it also enables ROCK to target Thr-853. As shown in Table 1, ROCK clearly shows greater specificity for Thr-853 on MYPT relative to ZIPK. Phosphorylation of Thr-853 by ROCK has been implicated by Cohen and co-workers (6) in the regulation of MYPT binding to myosin. Therefore, co-localization of MYPT, ZIPK, and ROCK at the membrane would suggest a model in which ZIPK directly regulates MYPT activity through phosphorylation of Thr-696, whereas ROCK regulates interactions with myosin via Thr-853 phosphorylation.


Figure 7
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  		FIGURE 7.
Pathways affecting actin cytoskeleton arrangement. A, Rho-binding proteins, including ROCK and mDia, are recruited to cell membranes for activation. ROCK phosphorylates a number of effectors (ZIPK, MYPT1, and LIM kinase (LIMK)) to stimulate cytoskeletal rearrangement. B, transient transfection of activated forms of ROCK and Rho differentially induce stress fiber formation. ROCK{Delta} directs focused stress fiber patterns, whereas RhoA 63L generates parallel arrays. But coexpression of ROCK{Delta} with ZIPK modifies stress fibers from a focused to a parallel pattern reminiscent of Rho-induced stress fibers. Coexpression of ROCK{Delta} and mDia{Delta}N3 produces a similar parallel pattern (44).

 
In this study, in non-muscle cells we demonstrate that ZIPK can alter ROCK{Delta}3-induced stress fiber formation and that this effect is dependent upon an intact Thr-265 phosphorylation site in non-muscle cells. ROCK{Delta}3-induced stress fibers were highly focused, perhaps due to lack of the Rho-binding domain and subsequent cellular mislocalization, but coexpression of wild-type ZIPK resulted in widely distributed parallel stress fibers similar to those induced by Rho overexpression. A recent study showed that Rho-like parallel stress fibers were similarly observed upon coexpression of ROCK{Delta}3 and mDia, another Rho effector protein (44). Taken with our results, these data suggest that parallel pathways exist downstream of Rho that culminate in actin cytoskeleton rearrangement (Fig. 7A). ZIPK functions downstream of ROCK{Delta}3 in one such pathway perhaps to direct stress fiber formation signals to the entire cell rather than to focused points. In this scenario, ZIPK acts as a soluble ROCK effector to signal outward from the cell membrane to the actomyosin cytoskeletal elements within the cytoplasm (Fig. 7B). This model is appealing because it would resolve the spatial discrepancy between ROCK and cytosolic actin fibers by placing ZIPK as a signaling intermediary during smooth muscle contraction or non-muscle cell motility (17, 28). Alternatively overexpressed ZIPK may compete with other substrates to abrogate ROCK effects on the actin cytoskeleton. Either mode of action must depend upon an intact Thr-265 site because ZIPK T265A coexpression resulted in little effect on ROCK{Delta}3-induced actin rearrangements.

Our studies now raise the questions: do all pathways governing Ca2+-independent phosphorylation of myosin in smooth muscle proceed through ZIPK, and does this mechanism extend to all non-muscle cells? Until recently, neither ROCK nor Rho has necessarily been thought of as a major initiator of non-apoptotic cell death in the manner of ZIPK or other members of the death-associated protein kinase family (35). However, Olson and co-workers (53) have provided evidence that ROCK participates in caspase-dependent apoptosis. Caspases cleave ROCK to produce constitutive activation, which in turn induces actin-myosin-driven nuclear disintegration, an effect that is blocked by Y-27632 (54-56). Until data shown herein, no reports to date have directly identified Rho/ROCK in the activation of ZIPK despite the extensive literature currently surrounding both kinases.

If ROCK is not the physiological regulator of ZIPK in mediating non-apoptotic cell death, what other factors may signal to ZIPK to initiate death responses? At least one group has suggested that the closely related kinase, DAPK1, is an immediate upstream activator of ZIPK. In this study, DAPK1 and various truncation mutants of DAPK1 were coexpressed in HEK293 with a ZIPK K42A mutant. This resulted in phosphorylation of ZIPK at Thr-229 and five other inferred sites (using block mutagenesis) between Ser-309 and Ser-326 (36). In our hands phosphorylation at Thr-299 is not activating but results in increased localization to the cytosol and potentially to myosin. Other than Ser-311, as reported in Graves et al. (34), we have not found any direct evidence for phosphorylation in vivo or in vitro between 308 and 326. Additionally we would question whether DAPK is in fact a physiological relevant ZIP kinase kinase. In the described DAPK study, the ZIPK K42A mutant was used as the substrate for DAPK both in vitro and in cell transfection experiments (36). Substitution of this conserved lysine has long been known to only marginally affect the Kd for ATP and is generally not considered the mutation of choice for rendering most protein kinases fully inactive (38, 57). Mutation at the K42A position (Lys-72 in cAMP-dependent protein kinase) renders kinases apparently inactive in standard in vitro kinase assays where ATP is often used in the low 10-50 µM range, but such mutations do not render kinases fully inactive at physiological ATP concentrations (5-10 mM) found in cells. In our hands, the ZIPK K42A mutant has considerable kinase activity at ATP concentrations >100 µM compared with the D161A mutant. Thus, autophosphorylation of ZIPK K42A cannot be ruled out, calling into question the conclusion that ZIPK is a direct target for DAPK in vivo. Experimental issues aside, if DAPK1 is a ZIP kinase kinase in vivo, clearly any signals that promote its activation are likely to be Ca2+/calmodulin-mediated, and it would be interesting to reexamine the role of DAPK1 in the regulation of ZIPK within this context. At least one report has directly linked phosphorylation at ZIPK Thr-265 with initiation of cell death in response to interleukin 6 family members. Thr-265 phosphorylation was stimulated in response to leukocyte inhibitory factor, and Thr -> Ala mutation of this site blocked leukocyte inhibitory factor-induced STAT3 phosphorylation in HEK293T cells (37). This finding suggests that a death-initiating Thr-265 ZIP kinase kinase likely exists but remains to be identified.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grant R01 HL07895. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental data. Back

1 Both authors contributed equally to this work. Back

2 To whom correspondence should be addressed: Dept. of Pharmacology and Cancer Biology, Duke University Medical Center, C119 LSRC Research Dr., Durham, NC 27710. Tel.: 919-613-8606; Fax: 919-668-0977; E-mail: hayst001{at}mc.duke.edu.

3 The abbreviations used are: LC20, 20-kDa regulatory light chain; SMPP-1M, myosin light chain phosphatase; ZIPK, zipper-interacting protein kinase; ROCK, Rho kinase; GTP{gamma}S, guanosine 5'-3-O-(thio)triphosphate; HEK, human embryonic kidney; MALDI, matrix-assisted laser desorption ionization; TOF, time-of-flight; HPLC, high pressure liquid chromatography; CRP, cleaved radioactive protein; MS, mass spectrometry; DAPK, death-associated protein kinase; STAT, signal transducers and activators of transcription; TERA, transitional endoplasmic reticulum ATPase; GEPH, Gephyrin; TCP, T-complex protein 1; PCCB, propionyl-CoA carboxylasebeta chain; ACTH, actin, {gamma}-enteric smooth muscle. Back


   ACKNOWLEDGMENTS
 
We thank Elizabeth Snyder for help with the preparation of figures and this manuscript.



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 ABSTRACT
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 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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