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Blood, 15 October 2008, Vol. 112, No. 8, pp. 3242-3254. Prepublished online as a Blood First Edition Paper on July 22, 2008; DOI 10.1182/blood-2007-12-126433. This Article Abstract Full Text (PDF) Supplemental Figures All Versions of this Article: blood-2007-12-126433v1 112/8/3242    most recent Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Rights and Permissions Citing Articles Citing Articles via CrossRef Citing Articles via Google Scholar Google Scholar Articles by Abair, T. D. Articles by Pozzi, A. Search for Related Content PubMed PubMed Citation Articles by Abair, T. D. Articles by Pozzi, A. Related Collections Hemostasis, Thrombosis, and Vascular Biology Social Bookmarking                   What's this?  Previous Article  |  Table of Contents  |  Next Article  HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY Functional analysis of the cytoplasmic domain of the integrin 1 subunit in endothelial cells Tristin D. Abair1,2, Nada Bulus1, Corina Borza1, Munirathinam Sundaramoorthy1, Roy Zent14, and Ambra Pozzi1,2,4 Departments of1 Medicine (Division of Nephrology), 2 Cancer Biology, and 3 Cell and Developmental Biology, Vanderbilt University, Nashville, TN; and 4 Department of Medicine, Veterans Affairs Hospital, Nashville, TN    Abstract Top Abstract Introduction Methods Results Discussion Authorship References   Integrin 1β1, the major collagen type IV receptor, is expressed by endothelial cells and plays a role in both physiologic and pathologic angiogenesis. Because the molecular mechanisms whereby this collagen IV receptor mediates endothelial cell functions are poorly understood, truncation and point mutants of the integrin 1 subunit cytoplasmic tail (amino acids 1137-1151) were generated and expressed into 1-null endothelial cells. We show that 1-null endothelial cells expressing the 1 subunit, which lacks the entire cytoplasmic tail (mutant 1-1136) or expresses all the amino acids up to the highly conserved GFFKR motif (mutant 1-1143), have a similar phenotype to parental 1-null cells. Pro1144 and Leu1145 were shown to be necessary for 1β1-mediated endothelial cell proliferation; Lys1146 for adhesion, migration, and tubulogenesis and Lys1147 for tubulogenesis. Integrin 1β1–dependent endothelial cell proliferation is primarily mediated by ERK activation, whereas migration and tubulogenesis require both p38 MAPK and PI3K/Akt activation. Thus, distinct amino acids distal to the GFFKR motif of the 1 integrin cytoplasmic tail mediate activation of selective downstream signaling pathways and specific endothelial cell functions.    Introduction Top Abstract Introduction Methods Results Discussion Authorship References   Angiogenesis, the formation of new blood vessels from preexisting vessels, is required for both physiologic and pathologic events, including embryonic development, wound healing, and tumor growth.1,2 Angiogenesis is a multistep process that requires endothelial cell proliferation, migration, adhesion to the vessel basement membrane, and formation of cell-cell junctions. Cell-matrix interactions, which are required for most of these cellular processes, are primarily mediated by integrins, transmembrane receptors for extracellular matrix components.3 Several integrin family members, including vβ3, vβ5, 5β1, 1β1, and 2β1, are expressed on endothelial cells and play a role in angiogenesis.4–10 The best studied are the RGD binding v and 5β1 integrins, and their role in angiogenesis is controversial, as they can be both proangiogenic and antiangiogenic.4,6,11,12 The function of the 2 major collagen binding receptors, integrins 1β1 and 2β1, in the control of endothelial cell functions is less clearly defined. Integrin 1β1 is thought to be proangiogenic as functional blocking antibodies or targeted deletion of the 1 subunit results in a decrease of both vascular endothelial growth factor-mediated and tumor-associated angiogenesis.7,8,13–15 In contrast, integrin 2β1 is proposed to be antiangiogenic, as deletion of the 2 subunit results in increased wound- and tumor-associated vasculature.9,16 The mechanism whereby integrin 1β1 promotes increased angiogenesis is poorly understood. We previously showed that integrin 1–null mice have smaller and less vascularized tumors than their wild-type counterparts. This effect is due in part to increased levels of circulating matrix metalloproteinase-9 in the 1-null mice, which results in increased generation of angiostatin, a potent inhibitor of endothelial cell proliferation, from circulating plasminogen.7,8,14,15 In addition, integrin 1β1 is known to promote cell survival and proliferation on collagenous substrata (key components of the vascular basement membrane) via activation of the Shc/Grb2/ERK pathway,17 suggesting that impaired integrin 1–dependent intracellular signaling in endothelial cells may also contribute to the abnormalities found in integrin 1–null mice. The extracellular domain of the integrin subunits is responsible for the specificity of ligand binding, whereas the cytoplasmic and transmembrane domains are important in regulating integrin activation and signaling.18–22 The activation state of integrins is thought to be dependent on interactions between the and β integrin tails. The highly conserved GFFKR motif in the tails is proposed to form a salt bridge with a highly conserved sequence HDRRE in the juxtamembrane region of the β tail. This interaction holds the integrin in the inactive state characterized by low ligand-binding affinity.23–27 On binding of intracellular proteins, such as talin or kindlins, to the β tail the juxtamembrane β tail interaction is thought to be disrupted, resulting in integrin activation and increased ligand-binding affinity.26,28–33 This mechanism has been primarily studied for the highly modulatable β2 and β3 integrins, where membrane proximal deletions of the GFFKR motif results in a constitutively activated integrin.23,34,35 The role of the GFFKR motif in modulating the activation state of β1-containing integrin is poorly defined. The cytoplasmic tail is also critical in mediating "outside-in" integrin signaling by interacting with adaptor, cytoskeletal, and signaling molecules.36–39 The importance of the 1 cytoplasmic tail in 1β1 integrin function is demonstrated by the requirement of this domain for cell spreading as well as focal adhesion and stress fiber formation.40,41 The 1 tail has also been shown to interact either directly or indirectly with several signaling molecules, including Shc, T-cell protein tyrosine phosphatase, phospholipase C, FAK, and PRL-3.37,38,40–42 However, little is known about the specific region(s) within the 15 amino acids of the 1 subunit cytoplasmic tail that control cellular function and signaling. To define which domains of the 1 cytoplasmic tail are required to mediate integrin 1β1–dependent endothelial cell function, truncation and point mutants within the tail were generated and expressed into integrin 1–null endothelial cells. Using these cells, we show that parental 1-null cells, 1-null cells expressing mutants lacking the entire cytoplasmic tail (mutant 1-1136), or expressing all the amino acids up to the GFFKR motif (mutant 1-1143) have a similar phenotype characterized by reduced adhesion, migration, proliferation, and tubulogenic potential on collagen substrata. In addition, we identify the specific residues distal to the GFFKR motif that control different integrin 1β1–dependent cell functions and activation of selective downstream signaling pathways.    Methods Top Abstract Introduction Methods Results Discussion Authorship References   Generation of mutant integrin 1 subunits Full-length human integrin 1 cDNA in the pLEN vector was a generous gift from Dr E. Marcantonio (Columbia University, New York, NY). The 1 subunit truncation mutants were generated by polymerase chain reaction, using the mature full-length 1 as a template.43 A common sense primer 5'-CGGTACCACCATGTTCAATGTTGATGTGAAAAACTC-3', containing a Kpn I restriction site and an ATG start codon, was used for all the truncation constructs. The antisense primers containing a Kpn I restriction site and stop codon were as follows: 5'-TAATGGTACCTCATCATTTCTCCATTTTCTTTTTCAG-3' (full-length 1); 5'-TAATGGTACCTCATCACTCCTCCATTTTCTTTTTCAG-3' (1-K1151E); 5'-TAATGGTACCTCATCACTCCATTTTCTTTTTCAGTGGTC-3' (mutant 1–1150); 5'-TAATGGTACCTCATCATTTCATTTTCTTTTTCAGTGGTC-3' (mutant 1-E1150K), 5'-TAATGGTACCTCATCACATTTTCTTTTTCAGTGGTCTTTTG-3' (mutant 1-1149); 5'-TAATGGTACCTCATCATTTCTT-TTTCAGTGGTCTTTTG-3' (mutant 1-1148), 5'-TAATGGTACCTCAT-CACAGTGGTCTTTTGAAGAATCC-3' (mutant 1-1145); 5'-TAATGG-TACCTCATCATCTTTTGAAGAATCCAATCTTCC-3' (mutant 1-1143); 5'-TAATGGTACCTCATCACCACAGTGCTAAAATGAGCAG-3' (mutant 1-1136). The single point mutants K1146A, K1147A, and K1148A were generated using the QuikChange II XL Site-Directed Mutagenesis Kit (Stratagene, La Jolla, CA) with appropriate primers. All mutants were checked by sequencing to verify they were correct. The various constructs were subcloned into the LZRS-GFP vector, modified from the original LZRS vector,44 to allow bicistronic expression of the protein of interest and GFP (a generous gift from Dr A. Reynolds).45 These constructs were transfected into the packaging cell line Phoenix 293 using FuGENE 6 transfection reagent according to the manufacturer's instructions (Roche Diagnostics, Indianapolis, IN). Cell culture and generation of cell populations Integrin 1–null pulmonary microvascular endothelial cells were isolated from integrin 1–null mice crossed with the immorto-mouse background, as described.7 Cells were propagated at 33°C in microvascular endothelial-cell medium-2 (EGM-2 MV) containing 5% fetal calf serum (Lonza Walkersville, Walkersville, MD) in the presence of 100 IU/mL of -interferon. For experiments, cells were cultured at 37°C in EGM-2 MV medium without -interferon for at least 4 days before use as this is the optimal time for the immorto endothelial cells to acquire a phenotype similar to freshly isolated primary endothelial cells. For the generation of 1-null endothelial cells expressing the various integrin 1 mutants, cells were infected for 2 weeks with virus contained in the medium of transfected Phoenix 293 cells. In some experiments, Chinese Hamster Ovary (CHO) cells were stably transfected with PCDNA3.1 or full-length integrin 1, 1-E1150K, and 1-K1151E subcloned into PCDNA3.1. Cells were transfected using Lipofectamine and Plus reagent (Invitrogen, Carlsbad, CA) followed by zeocin selection (Sigma-Aldrich, St Louis, MO) according to the manufacturer's instructions. Stable cell populations of 1-null endothelial cells or CHO cells expressing equal levels of the various mutants were selected by flow cytometry using antibodies recognizing the extracellular I domain of human integrin 1 (MAB1973; Chemicon International, Temecula, CA). Polyclonal cell populations were used to avoid some of the pitfalls associated with isolation of monoclonal cell lines. Cell morphology Cells were plated in serum-free medium on chamber slides coated with 10 µg/mL fibronectin (a non–integrin 1β1 ligand, used as positive control) or collagen IV (a specific integrin 1β1 ligand; both from Sigma-Aldrich). After 4 hours, the cells that remained adherent were fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton X-100, and stained with rhodamine-phalloidin (Invitrogen). Cells were subsequently washed with phosphate-buffered saline (PBS) and examined under a fluorescence microscope (Nikon, Tokyo, Japan). Three independent experiments were performed. Cell adhesion The cell adhesion assay was performed as previously described.46 Briefly, 96-well plates were coated with fibronectin (10 µg/mL) or collagen IV (at various concentrations) for 1 hour at 37°C. After blocking nonspecific adhesion with 1% bovine serum albumin in PBS, 5 x 104 cells in 100 µL serum-free medium were added to the plates and incubated for 1 hour at 37°C. In some experiments, cells were preincubated with 10 µM PD169316 (a p38 MAPK inhibitor), 10 µM PD98059 (a MEK inhibitor), or 5 µM LY294002 (a PI3K inhibitor; all from Calbiochem) for 30 minutes before the assay. After removing nonadherent cells, the cells were fixed with 4% formaldehyde, stained with 1% crystal violet, solubilized in 20% acetic acid, and the absorbance read at 595 nm. Cell adhesion to 1% bovine serum albumin–coated wells was subtracted from the values obtained on ECM proteins. Four independent experiments were performed in quadruplicates. Cell migration Cell migration was assayed in transwells consisting of polyvinylpyrolidone-free polycarbonate filters with 8-µm pores (Costar; Corning, Cambridge, MA) as previously described.46 The bottoms of the filters were coated with collagen IV or fibronectin (10 µg/mL in PBS), and 105 cells were used for each assay. In some experiments, 5 µM LY294002, 10 µM PD98059, or 10 µM PD169316 were added to both top and bottom wells. Cells were allowed to migrate for either 6 (CHO cells) or 16 (endothelial cells) hours at 37°C, after which they were fixed in 4% formaldehyde, stained with 1% crystal violet, and 5 randomly chosen fields counted at 200x magnification. Four independent experiments were performed in duplicates. Tubulogenesis Capillary-like formation on collagen and fibrin gels was analyzed as described.46 For collagen gels, 50 µL gel composed of 1 mg/mL rat tail collagen I (BD Biosciences, San Jose, CA), 30 µg/mL collagen IV, and Dulbecco minimal essential media containing 20 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (pH 7.2) were allowed to polymerize in 96-well plates. For fibrin gels, fibrinogen (2.5 mg/mL) was dissolved in PBS and clotting was started by the addition of thrombin (0.250 U/mL); 50 µL of the mixture was added to 96-well plates and allowed to polymerize for 10 minutes at 37°C. Endothelial cells (1.5 x 104 in 150 µL serum-free medium) were then plated on solidified gels in the presence or absence of the kinase inhibitors indicated in "Cell adhesion." After 12 hours, the cells were fixed with 4% formaldehyde in PBS, and phase-contrast photomicrographs were taken using a Nikon inverted microscope. To quantify capillary-linked network formation, cellular nodes were defined as junctions linking at least 3 cells, and they were counted from digital images. Four independent experiments were performed with a total of 20 images analyzed per cell population. Cell proliferation Endothelial cells (5 x 103/well) or CHO cells (103/well) were plated in low serum (1% fetal calf serum) onto 96-well plates coated with 10 µg/mL collagen IV or 10 µg/mL fibronectin. Under these plating conditions, all cell populations, independent of the genotype or construct expressed, adhere and spread on both matrices. After 4 hours, the cells were gently washed and incubated with serum-free medium (thus allowing integrin-mediated signaling to control cell proliferation) with or without 5 µM LY294002, 10 µM PD98059, or 10 µM PD169316 in the presence of [3H]thymidine (0.5 µCi/well). After 48 hours, the cells were collected and the amount of incorporated [3H]thymidine analyzed as previously described.7 Four independent experiments were performed in quadruplicates. Western blot analysis To determine the activation of intracellular downstream signaling, serum-starved endothelial cells were embedded for different amount of times in 30 µL collagen I + IV gels, prepared as described in "Tubulogenesis" (3.5 x 104 cells/gel); or plated for different amount of times onto dishes coated with 10 µg/mL fibronectin (5 x 105 cells/10-cm dishes). In some experiments, human anti–integrin 1 or mouse anti–integrin 2 antibodies were used at the final concentration of 30 µg/mL. For cells in the gels, equal volumes of Laemmli buffer containing β-mercaptoethanol were then added to the gels. Samples were then sonicated, boiled, and run onto a 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) gel and subsequently transferred to nitrocellulose membranes. For cells on fibronectin, adherent cells were scraped and lysed, and equal amounts of total cell lysates (20 µg/lane) were run onto a 10% SDS-PAGE. Membranes were incubated with antiphospho-ERK, antiphospho-p38 MAPK, antiphospho-Akt, anti-ERK, anti-p38 MAPK, or anti-Akt antibodies (all from Cell Signaling Technology, Danvers, MA) followed by the appropriate horseradish peroxidase-conjugated secondary antibodies. Immunoreactive bands were identified using enhanced chemiluminescence according to the manufacturer's instructions. Four independent experiments were performed. Phosphorylated and total ERK, p38 MAPK, and Akt bands were quantified by densitometry analysis, and the levels of phosphorylated signal were expressed as phosphorylated kinase/total kinase ratio. Values were expressed as the mean plus or minus SD of 3 experiments. Immunoprecipitation assay The different endothelial cell populations were lysed in a lysis buffer consisting of 50 mM Tris-HCl (pH 7.5), 10 mM MgCl2, 200 mM NaCl, 5% glycerol, 1% NP-40, 0.05% Tween-20, 1 mM NaV, 1 mM NaF, and 1x proteinase inhibitor. The lysates were centrifuged at 12 000g for 10 minutes, and the supernatants were used for immunoprecipitation; 1 mg total cell lysates were precleaned with 20 µL packed protein-G Sepharose beads (GE Healthcare, Little Chalfont, United Kingdom) and subsequently incubated with 5 µg anti–human integrin 1 antibodies (TS 2/7; Santa Cruz Biotechnology, Santa Cruz, CA) and 30 µL packed protein-G Sepharose and incubated overnight at 4°C. Beads were washed 3 times with lysis buffer, and bound proteins were eluted by boiling the beads in SDS-PAGE sample buffer under reducing conditions. Samples were separated onto a 10% SDS gel and subsequently transferred to nitrocellulose membranes. Membranes were then cut at the level of the 150-kDa marker. The upper part of the membranes was incubated with an antihuman integrin 1 antibody (MAB1973; Chemicon International), whereas the lower part of the membranes was incubated with an antimouse integrin β1 antibody (Chemicon International) followed by the appropriate horseradish peroxidase-conjugated secondary antibodies; 40 µg total cell lysates was analyzed for ERK levels to verify loading. Statistical analysis The Student t test was used for comparisons between 2 groups, and analysis of variance using Sigma-Stat software for statistical difference between multiple groups. A value of P less than .05 was considered statistically significant.    Results Top Abstract Introduction Methods Results Discussion Authorship References   Specific amino acids of the integrin 1 tail mediate endothelial cell spreading, adhesion, migration, proliferation, and tubulogenesis To identify the amino acids in the 1 integrin subunit cytoplasmic domain that regulate endothelial cell proliferation, migration, and tubulogenesis, truncation mutants of the human integrin 1 cDNA were generated (Figure 1A). Full-length or mutated human 1 integrin subunits were expressed into mouse 1-null (1KO) endothelial cells, and cell populations expressing similar levels of this subunit were isolated by fluorescence-activated cell sorter (FACS; Figure 1B). Formation of chimeric dimers between the human integrin 1 and the mouse β1 subunits was confirmed by immunoprecipitation assays (Figure 1C). Thus, the human integrin 1 subunit forms a complex with the mouse β1 subunit, as previously suggested for other human subunits expressed in mouse cells.46–48 View larger version (25K): [in this window] [in a new window]   Figure 1. Amino acids 1146-1148 of the integrin 1 tail control cell spreading, adhesion, and migration. (A) Schematic representation of the cytoplasmic truncation mutants of the human integrin 1 cDNA. (B) 1-null endothelial cells were transduced with either empty vector (1KO) or the human integrin 1 mutant cDNAs indicated in panel A, and cell populations with equal levels of expression of the integrin 1 subunits were sorted by FACS using anti–human integrin 1 antibodies. (C) One millligram of total cell lysates of the cell populations indicated was immunoprecipitated with anti–human integrin 1 antibodies, subjected to SDS-PAGE, and immunoblotted with antibodies to either human integrin 1 or mouse β1 subunits. Equal loading was confirmed by analyzing the levels of total ERK in 40 µg total cell lysates. (D,E) Integrin 1KO endothelial cells expressing either the empty vector or cytoplasmic tail deletion mutants were plated in serum-free medium on 10 µg/mL collagen IV (D) or fibronectin (E). After 4 hours, the cells were fixed and stained with rhodamine-phalloidin. A representative cell is shown for each cell population. Bar represents 10 µm. (F,G) The cell populations were plated in serum-free medium on collagen IV at the concentrations indicated (F) or 10 µg/mL collagen IV with or without anti–human integrin 1 antibodies (10 µg/mL) (G) for 1 hour and their adhesion determined as described in "Cell adhesion." Values are mean plus or minus SD of one representative experiment performed in quadruplicate. (H,I) The cell populations were plated on serum-free medium transwells coated with 10 µg/mL collagen IV with () or without () anti–human integrin 1 antibodies (10 µg/mL), and migration was evaluated 16 hours after plating. Values are mean plus or minus SD of one representative experiment performed in duplicate (5 fields/transwell were analyzed). Differences between 1KO and 1 mutant–expressing cells (*) and antibody-untreated versus-treated 1 mutant–expressing cells (**) were significant with P < .05.   To investigate functionality of the mutants, cells were plated in serum-free medium on either collagen IV (the major integrin 1β1 binding ligand) or fibronectin (an integrin 1β1–independent ligand) and their morphology evaluated after 4 hours. As previously described for fibroblasts,49 approximately 5% to 10% of the 1KO endothelial cells adhered to the collagen IV under these conditions but failed to spread (Figure 1D). This phenotype was reversed by expression of the full-length 1 subunit (Figure 1D). Like the 1KO cells, approximately 5% to 10% of the of the endothelial cells expressing the 1-1136, 1-1143, or 1-1145 mutants adhered to collagen IV but failed to spread (Figure 1D). In contrast, endothelial cells transfected with the 1-1148 mutant spread to a similar extent as cells reconstituted with full-length 1 (Figure 1D). No differences in spreading among the different mutants were observed on fibronectin (Figure 1E), confirming that the phenotypes observed were the result of alterations in integrin 1β1 interactions with collagen IV. Similar to cell spreading, only cells transfected with either full-length 1 or the 1-1148 mutants adhered in a dose-dependent manner and migrated on collagen IV (Figure 1F,H). Both cell adhesion and migration on collagen IV were inhibited by antihuman integrin 1 antibodies, proving these cell functions were integrin 1β1 dependent (Figure 1G,I). Finally, only cells expressing either full-length 1 or the 1-1148 mutants underwent tubulogenesis on collagen I + IV gels (Figure 2A), and this event was inhibited by antihuman integrin 1 antibodies (not shown). Interestingly, both the 1-1145– and 1-1148–expressing cells proliferated on collagen IV to the same degree as cells reconstituted with full-length 1 (Figure 2B). Thus, Pro1144 and Leu1145 within the 1 cytoplasmic tail are required for proliferation, whereas the triple lysine motif (amino acids 1146-1148) is required for 1β1-dependent cell spreading, adhesion, migration, and tubulogenesis on collagen IV. View larger version (29K): [in this window] [in a new window]   Figure 2. Integrin 1 cytoplasmic tail mutants activate distinct signaling pathways in endothelial cells. (A) The cell populations were plated in serum-free medium on solidified collagen I + IV gels. Tubulogenesis was quantified 16 hours after plating as described in "Tubulogenesis." Values are mean plus or minus SD of 4 independent experiments. Gels were viewed with a Nikon Diaphot inverted research microscope (Nikon, Tokyo, Japan) using a lens at 20x/0.2 Ph2 LD 0.4. Images were acquired using a Canon PowerShot S5 IS camera (Canon USA, Lake Success, NY) and were processed with Adobe Photoshop version 9.0 software (Adobe Systems, San Jose, CA). (B) The cell populations were plated in 96-well plates coated with 10 µg/mL collagen IV. Four hours later, the cells were incubated with serum-free medium containing 3H-thymidine (0.5 µCi/well) for a further 48 hours, and proliferation was then evaluated as described in "Cell proliferation." Values are mean plus or minus SD of one representative experiment performed in quadruplicate. *Statistically significant differences (P < .05) between the 1KO cells and 1 mutant–expressing cells. (C) The cell populations were serum-starved for 24 hours and embedded in collagen I + IV gels for the time indicated. The gels were sonicated and run on an SDS-PAGE gel to detect levels of activated and total p38 MAPK, ERK, and Akt. Images are representative of 3 independent experiments. Vertical line(s) have been inserted to indicate a repositioned gel lane. (D) Phosphorylated and total kinase bands were quantified by densitometry analysis, and the phosphorylated signal was expressed as phosphorylated kinase/total kinase ratio. Values are the mean plus orf minus SD of 3 independent experiments. * indicates significant differences (P < .05) relative to 1KO cells. (E-G) The cell populations indicated were subjected to migration (E), tubulogenesis (F), and proliferation (G) assays on collagen IV or fibronectin in the presence or absence of 10 µM PD169316, 10 µM PD98059, or 5 µM LY294002. Values are the mean plus or minus SD of one representative experiment. Differences between untreated 1KO and 1 mutant–expressing cells (*) and inhibitor untreated versus treated 1 mutant–expressing cells (#) were significant with P < .05.   Distinct cellular functions are mediated by activation of different 1 integrin subunit-dependent signaling pathways Although we previously showed that integrin 1β1 confers the ability of fibroblasts to proliferate on collagenous substrata via activation of the Shc/Grb2/ERK pathway,17 little is known about other pathways regulated by this collagen receptor. Because p38 MAPK, ERK, and Akt activation is required for the regulation of different endothelial cell functions,50 the various mutant expressing cells were embedded in collagen I + IV gels and activation of these 3 kinases analyzed at different time points. Significant basal and collagen I + IV-mediated p38 MAPK activation was only seen in the 1–1148- and full-length 1-expressing cells (Figure 2C,D), and collagen I + IV–induced Akt phosphorylation was detected in the same cell populations (Figure 2C,D). Interestingly, collagen I + IV–mediated ERK activation was seen in the 1-1145–, 1-1148–, and full-length 1-expressing cells (Figure 2C,D). When 1KO and full-length 1-expressing endothelial cells were plated on fibronectin, both cell types activated all 3 kinases equally within 10 minutes of plating (Figure S1Figure S1A,B, available on the Blood website; see the Supplemental Materials link at the top of the online article), indicating that loss and/or reexpression of the integrin 1 subunit only affects collagen-mediated signaling. Altogether, these results suggest that, in a collagenous milieu, ERK activation appears to correlate with mutants able to support cell proliferation (Figure 2B), whereas activation of p38 MAPK and Akt correlated with mutants able to support migration and tubulogenesis (Figures 1H, 2A). To determine whether integrin 1β1–mediated signaling correlated with the ability of this receptor to induce cell migration, tubulogenesis, and proliferation, these assays were performed in the presence of selective kinase inhibitors. The p38 MAPK inhibitor significantly decreased the ability of cells expressing 1-1148 and full-length 1 to migrate on collagen IV, whereas the MEK and PI3K inhibitors had no effect (Figure 2E). When migration on fibronectin was analyzed, 1KO- and full-length 1-expressing endothelial cells showed comparable migration rates, which was inhibited to a similar extent by the p38 MAPK inhibitor (Figure 2E). Both the p38 MAPK and PI3K inhibitors inhibited the ability of the 1-1148 and full-length 1-expressing cells to form tubes on collagen I + IV gels (Figure 2F). When 1KO- and full-length 1-expressing endothelial cells were plated onto fibrin gels, both cell types underwent tubulogenesis that was equally inhibited by the p38 MAPK and PI3K inhibitors (Figure S1Figure S1C,D). Proliferation of the 1-1145–, 1-1148–, and full-length 1-expressing cells on collagen IV was predominantly inhibited by the MEK inhibitor and, to a lesser extent, by the PI3K inhibitor (Figure 2G). In contrast, proliferation of the 1KO- and full-length 1-expressing endothelial cells on fibronectin was inhibited to a greater extent than cells grown on collagen IV by both MEK and PI3K inhibitors (Figure 2G). These results suggest that (1) integrin-dependent migration of endothelial cells, including that mediated by 1β1, requires the p38 MAPK pathway; (2) the p38 MAPK and PI3K signaling pathways are required for both 1β1-dependent and -independent tubulogenesis; (3) ERK, and to a lesser extent PI3K, plays a role in integrin 1β1-mediated proliferation; and (4) both the ERK and PI3K pathways play an equally important role in regulating endothelial cell proliferation on fibronectin. Lys1146 is required for integrin 1β1-dependent endothelial cell adhesion, migration and tubulogenesis, whereas Lys1147 is only required for tubule formation The 6 COOH-terminal amino acids of the integrin 1 cytoplasmic tail (KKKMEK) were required for cell spreading, adhesion, migration, and tubulogenesis (Figures 1D-I, 2A); however, the last 3 amino acids (MEK) were dispensable, allowing us to hypothesize that Lys1146-1148 were required to mediate these cell functions. For this reason, we mutated each lysine individually as well as all 3 lysines to alanines (Figure 3A) and attempted to express the mutants into 1KO endothelial cells. Whereas generation of cell populations expressing the single mutants was successful (Figure 3B,C), we were unable to obtain surface expression of the triple mutant. The K1146A-expressing cells failed to adhere, migrate, or undergo tubulogenesis on collagen IV (Figures 3D,F, 4A). In contrast, the K1147A mutant cells adhered and migrated to a similar extent as endothelial cells reconstituted with full-length 1 (Figure 3D,F) but were unable to undergo tubulogenesis on collagen IV (Figure 4A). No differences in adhesion, migration, and tubulogenesis were observed between the K1148A and full-length 1-expressing cells (Figures 3D,F, 4A). The observation that adhesion and migration of K1147A and K1148A mutant cells were inhibited by antihuman integrin 1 antibodies (Figure 3E,G) confirmed that these effects were indeed integrin 1β1–dependent. View larger version (25K): [in this window] [in a new window]   Figure 3. K1146A point mutation of the integrin 1 cytoplasmic tail leads to decreased endothelial cell adhesion and migration. (A) Schematic representation of single point mutants (K/A) generated from the full-length integrin 1 subunit. (B) 1KO endothelial cells were transduced with either empty vector or the integrin 1 mutant cDNAs indicated in panel A, and cell populations were sorted by FACS using antihuman integrin 1 antibodies. (C) Total cell lysates of the cell populations were used to detect the levels of full-length and mutant human integrin 1 as well as mouse β1 subunits as described in detail in Figure 1C. (D-G) Cell adhesion on different concentrations of collagen IV (D) or in the presence of anti–human integrin 1 antibodies (E) as well as migration in the absence (F) and presence (G) of anti–human integrin 1 antibodies were determined as described in Figure 1. Note that only cells expressing K1146A mutant fail to adhere and migrate on collagen IV. * indicates statistically significant differences (P < .05) between the 1KO and the 1 mutant–expressing endothelial cells; * indicates statistically significant differences (P < .05) between untreated and antibody-treated 1 mutant–expressing cells. (H,I) The endothelial cells indicated were plated on 10 µg/mL collagen IV or fibronectin. After 4 hours, the cells were fixed and stained with rhodamine-phalloidin. A representative cell is shown for each cell population. Bar represents 10 µm.   View larger version (30K): [in this window] [in a new window]   Figure 4. K1146A and K1147A point mutations of the integrin 1 cytoplasmic tail lead to decreased tubulogenesis. Tubulogenesis (A), cell proliferation (B), cell signaling (C), and densitometry analysis (D) were determined as described in Figure 2. Images of cells undergoing tubulogenesis were taken as described in Figure 2A. *Statistically significant differences (P < .05) between the 1KO and the 1 mutant–expressing endothelial cells. Vertical line(s) in panel C have been inserted to indicate a repositioned gel lane.   When cell morphology was assessed, only 5% to 10% of the K1146A mutant cells adhered on collagen IV; however, these adherent cells spread in a manner similar to K1147A-, K1148A-, and full-length 1-expressing cells (Figure 3H). As expected, none of the single K/A substitutions affected cell spreading on fibronectin (Figure 3I). These results suggest that Lys1146 is necessary for cell adhesion, migration, and tubulogenesis, whereas Lys1147 is required primarily for tubulogenesis. Finally, the single mutant-expressing cells proliferated to levels observed in full-length 1-expressing cells (Figure 4B), supporting the finding that cell proliferation is primarily mediated by the Pro1144 and Lys1145 NH2 terminal to these 3 lysine residues. When the ability of the K/A point mutants to activate p38 MAPK, Akt, and ERK was determined, the K1147A-, K1148A-, and full-length 1-expressing cells activated p38 MAPK to the same degree both at baseline and after collagen stimulation (Figure 4C,D). Only the K1148A mutant cells activated Akt to the same degree as cells expressing full-length 1 after collagen stimulation (Figure 4C,D). ERK activation in all mutants was the same as full-length 1-expressing cells (Figure 4C,D). Thus, the ability of these mutants to activate p38 MAPK correlated with endothelial cell migration, whereas Akt activity correlated with the ability of endothelial cell to undergo tubule formation. Deletion of Lys1151 of the integrin 1 cytoplasmic tail impairs endothelial cell adhesion, migration, and tubulogenesis As endothelial cells expressing the 1-1148 mutant displayed comparable functions to cells expressing full-length 1, we hypothesized that deleting both Glu1150 and Lys1151 (mutant 1-1149) or only Lys1151 (mutant 1-1150) (Figure 5A) would not alter integrin 1β1–mediated function. Cell populations expressing comparable levels of both 1-1149 and 1-1150 mutants (Figure 5B,C) spread on collagen IV and fibronectin to a similar extent as 1-1148 or 1 full-length expressing cells within 4 hours from plating (Figure 5D,E). The 1-1149 cells also adhered, migrated, and formed tubules, such as 1-1148 or 1 full-length expressing cells (Figure 6A-C), and these events were integrin 1β1–dependent (not shown). Despite their ability to spread, the 1-1150 cells adhered, migrated, and formed tubules significantly less than 1-1148 or 1 full-length expressing cells (Figure 6A-C). No differences in cell proliferation were observed between these 2 mutants and 1 full-length expressing cells (Figure 6D). As expected, the 1-1149 cells activated p38 MAPK, ERK, and Akt in response to collagen stimulation, whereas the 1-1150 cells only activated ERK (Figure 6E,F). View larger version (23K): [in this window] [in a new window]   Figure 5. Generation of integrin 1-null endothelial cells expressing deletion mutants lacking Glu1150 and/or Lys1151 of the integrin 1 cytoplasmic tail. (A) Schematic representation of the cytoplasmic truncation mutants of the human integrin 1 cDNA. (B) 1KO endothelial cells were transduced with either empty vector or the integrin 1 mutant cDNAs indicated in panel A, and cell populations expressing the integrin 1 constructs were sorted by FACS using anti–human integrin 1 antibodies. (C) Total cell lysates of the cell populations were used to detect the levels of full-length and mutated human integrin 1 as well as mouse β1 subunits as described in detail in Figure 1C. (D,E) Integrin 1KO endothelial cells expressing vector alone or the truncation mutants were plated on 10 µg/mL collagen IV or fibronectin. After 4 hours, the cells were fixed and stained with rhodamine-phalloidin. A representative cell is shown for each cell population. Bar represents 10 µm.   View larger version (28K): [in this window] [in a new window]   Figure 6. Lys1151 within the integrin 1 tail is required for endothelial cell adhesion, migration, and tubulogenesis, but not proliferation. Cell adhesion (A), migration (B), tubulogenesis (C), and proliferation (D) were determined as described in Figures 1 and 2. (E,F) Integrin 1β1–dependent signaling and densitometry analysis were determined as described in Figure 2C,D. * indicates statistically significant differences (P < .05) between the 1KO and the 1 mutant–expressing endothelial cells.   Substitution of Lys1151 with a negatively charged amino acid impairs cell adhesion and migration These results suggest that the presence of a negatively charged amino acid at the COOH terminus of the integrin 1 tail impairs cell adhesion, migration, and tubulogenesis. To test this hypothesis, we generated populations of CHO cells expressing equal levels of the full-length integrin 1 subunit, the point mutant 1-K1151E, and the truncation mutant 1-E1150K (Figure 7A,B). CHO cells were used for this experiment because (1) making stable endothelial cell populations is extremely difficult and time-consuming; (2) similar to integrin 1–null endothelial cells, CHO cells show low baseline spreading, adhesion, migration, and proliferation on collagen IV (Figure 7)51; and (3) most importantly, they have been successfully used to analyze the role of the human integrin 1 subunit in cell functions.40,51 View larger version (21K): [in this window] [in a new window]   Figure 7. Negatively charged amino acids at the COOH-terminus of the integrin 1 tail inhibit cell adhesion and migration, but not proliferation. (A) Schematic representation of mutants generated from the full-length integrin 1 subunit. (B) CHO cells were transfected with either empty vector or the integrin 1 mutant cDNAs indicated in panel A, and cell populations were sorted by FACS using anti–human integrin 1 antibodies. (C) The CHO cell populations indicated were plated on 10 µg/mL collagen IV and after 1 hour they were fixed and stained with rhodamine-phalloidin. A representative cell is shown for each cell population. Bar represents 10 µm. (D-F) Cell adhesion (D), migration (E), and proliferation (F) were determined as described in Figures 1 and 2. * indicates statistically significant differences (P < .05) between the vector transfected and 1 mutant–expressing CHO cells.   As previously reported,51 vector-transfected CHO cells failed to spread on collagen IV, whereas CHO cells transfected with mutants 1-E1150K, 1-K1151E, or 1 full-length spread equally on this substrate (Figure 7C). The CHO cells expressing either full-length or the mutant 1-E1150K adhered and migrated on collagen IV significantly more than vector or 1-K1151E transfected CHO cells (Figure 7D,E). Finally, all the CHO cells expressing the different integrin 1 constructs proliferated significantly more than vector-transfected CHO cells on collagen IV (Figure 7F). These results strongly suggest that a positively charged amino acid at the COOH-terminus of the integrin 1 tail mediates integrin 1β1–dependent cell adhesion and migration.    Discussion Top Abstract Introduction Methods Results Discussion Authorship References   We previously showed that integrin 1β1 plays a role in pathologic angiogenesis,7,8,14,15 but the specific mechanism(s) by which the 1 subunit controls endothelial cell functions is unknown. Using a mutagenesis approach, we determined the critical residues within the 1 cytoplasmic tail, as well as the signaling pathways required for specific integrin 1β1–dependent endothelial cell functions. We show that: (1) mutants with deletion of the entire cytoplasmic tail (1-1136) or just distal to the GFFKR motif (1-1143) do not restore endothelial cell adhesion, migration, proliferation and tubulogenesis on collagen IV; (2) Pro1144 and Leu1145 are required for integrin 1β1–dependent endothelial cell proliferation; (3) Lys1146 is required for adhesion, migration, and tubulogenesis, whereas Lys1147 is only required for tubulogenesis; (4) a positively charged amino acid is most probably required at the COOH-terminus of the 1 tail to mediate cell adhesion, migration, and tubulogenesis; and (5) integrin 1β1–dependent cell migration requires p38 MAPK activation; tubulogenesis requires both p38 MAPK and PI3K activation, whereas proliferation is predominantly dependent on ERK activation. Thus, specific amino acids distal to the GFFKR motif are critical for activation of distinct signaling pathways and 1β1-dependent cell functions. The highly conserved GFFKR region in the juxtamembrane domain of the integrins has been proposed to bind to the β tail and keep the integrin in an inactive state.26,52 In this context, deletion of this region in the integrin IIb tail or mutations that disrupt integrin IIb-β3 interactions activates this receptor.26,52 In our study, 1KO endothelial cells expressing both 1-1136 (which lacks the GFFKR motif) and 1-1143 (which contains the GFFKR motif) have a similar phenotype to parental 1KO cells, suggesting that the highly conserved GFFKR motif might not play a role in regulating integrin 1 activation. This possibility is supported by the finding of Czuchra et al53 that mice carrying a D759A mutation in the integrin β1 tail HDRRE motif, which is thought to interact with the GFFKR of the subunits, had no obvious phenotype in vivo. Moreover, keratinocytes isolated from these mice showed normal adhesion, spreading, and migration in vitro.53 It is possible that the GFFKR in the 1 subunit does not play a role in integrin activation but is simply required for hetero- or homo-dimerization of the integrin and β subunits resulting in integrin clustering,54 which is critical for cell adhesion. This hypothesis is supported by the finding that CHO cells expressing an integrin 1 subunit lacking the cytoplasmic tail adhered but did not migrate on collagen IV, whereas a mutant containing the GFFKR motif induced cell migration and stress fiber formation.55 Although we attempted to perform binding assays in cell suspension to determine the role of different 1 mutants on 1β1 affinity for collagen IV, we were unable to identify a fragment that bound efficiently and specifically to the extracellular domain of this integrin. We show that the addition of Pro1144 and Leu1145 to the GFFKR motif is sufficient to support integrin 1β1–dependent endothelial cell proliferation and induce ERK activation. The inhibitor studies show this kinase is critical for 1β1-dependent cell proliferation. This result parallels our previous finding in fibroblasts that integrin 1β1 activates the Shc/Grb2/ERK pathway to promote proliferation on collagenous substrata.17 However, our observation that the first 9 amino acids of the 1 tail are required for ERK activation contrasts with the previous finding that the integrin 1 transmembrane, but not cytoplasmic domain, is required for integrin 1–mediated recruitment of Shc and consequent ERK activation.38 Our study suggests that a Shc-independent pathway might be responsible for integrin 1β1-mediated activation of ERK in endothelial cells and that Pro1144 and Leu1145 might serve as a binding site for an adaptor molecule(s) able to promote ERK activation in endothelial cells. Thus, it is possible that activation of ERK by integrin 1β1 is cell type–specific. We demonstrate that Lys1146 is required for cell adhesion, migration, and tubulogenesis, whereas Lys1147 is only required for tubulogenesis. These differences in cell function correlate with the finding that the K1146A mutation results in loss of both p38 MAPK and Akt activation, whereas the K1147A mutation only results in loss of Akt activation. Thus, these 2 lysines promote particular cell functions by activating specific signaling pathways. The observation that integrin 1–mediated p38 MAPK activation is required for endothelial cell migration and tubulogenesis agrees with the finding that activation of this kinase promotes endothelial cell migration.56,57 However, these data contrast with the findings that integrin 2 subunit is required for collagen-mediated p38 MAPK activation.47,58 The integrin 1KO endothelial cells (despite expressing endogenous integrin 2β1) cannot activate p38 MAPK when embedded in collagen gels, whereas reexpression of the integrin 1 subunit results in significant collagen-mediated p38 MAPK activation. Interestingly, incubation of full-length 1 reconstituted endothelial cells with either antihuman 1 or antimouse 2 antibodies significantly decreases the collagen-mediated p38 MAPK activation (Figure S2Figure S2A,B). Together, these data suggest that in endothelial cells both integrins 1β1 and 2β1 are required to promote collagen-mediated p38 MAPK activation, and blocking either integrin decreases p38 MAPK activation to the levels observed in 1KO cells. We demonstrate that integrin 1β1–mediated Akt activation plays a role in regulating endothelial cell tubulogenesis. The K1147A expressing cells, which cannot activate Akt or form tubules, show normal adhesion and migration, suggesting that tubulogenesis is regulated by cellular events other than cell migration. One possible mechanism might be cell polarization because PI3K inhibitors block cell polarization and directional migration of leukocytes and breast cancer cells.59,60 Moreover, inhibition of PI3K in endothelial cells prevents growth factor-induced branching or tubule formation without affecting other cell functions.61,62 One of the most surprising results in the study was that deletion of the terminal Lys1151 (mutant 1-1150) or substitution of this lysine with a glutamic acid (mutant 1-K1151E) resulted in the loss of integrin 1β1–dependent cell adhesion, migration, and tubulogenesis, whereas cells expressing mutant 1-1149 (where both Lys1151 and Glu1150 are deleted) or mutant 1-E1150K behaved like cells expressing full-length 1. A possible explanation for this result is that the presence of a positive or neutral (ie, 1-1150 mutant) charge at the COOH-terminus of the 1 tail determines how the tail is oriented relative to the β tail and/or to the plasma membrane, thus influencing the binding of signaling molecules to the COOH-terminus of the 1 cytoplasmic tail. In conclusion, we demonstrate that specific regions of the integrin 1 cytoplasmic tail are responsible for the activation of selective downstream signaling pathways that regulate specific endothelial cell functions. As integrin 1β1 plays a key role in angiogenesis, identification of the molecular mechanisms whereby this collagen binding receptor controls endothelial cell functions might have implications in determining whether this integrin is a good drug target for diseases characterized by uncontrolled new blood vessel formation.    Authorship Top Abstract Introduction Methods Results Discussion Authorship References   Contribution: T.D.A. performed experiments and made figures; N.B. performed experiments; C.B. contributed to the generation of reagents; M.S. contributed analytical tools and analyzed data; R.Z. analyzed data and wrote the paper; and A.P. designed the research and wrote the paper. Conflict-of-interest disclosure: The authors declare no competing financial interests. Correspondence: Ambra Pozzi, Department of Medicine, Division of Nephrology, Medical Center North, B3109, Vanderbilt University, Nashville, TN 37232; e-mail: ambra.pozzi{at}vanderbilt.edu.    Acknowledgments   The authors thank Cathy Alford at the Department of Veterans Affairs for help with the flow cytometric analysis. This work was supported by R01-CA94849 (A.P.), RO1-DK 69 921 and R01-DK 075594 (R.Z.), P01 DK65123 (R.Z., A.P.), and a Merit award from the Department of Veterans Affairs (R.Z.).    Footnotes   Submitted November 30, 2008; accepted June 26, 2008. Prepublished online as Blood First Edition Paper, July 22, 2008 DOI: 10.1182/blood-2007-12-126433 The online version of this article contains a data supplement. The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 USC section 1734.    References Top Abstract Introduction Methods Results Discussion Authorship References   Pandya NM, Dhalla NS, Santani DD. Angiogenesis: a new target for future therapy. Vascul Pharmacol. 2006;44:265–274.[CrossRef][Medline] [Order article via Infotrieve] Norrby K. In vivo models of angiogenesis. J Cell Mol Med. 2006;10:588–612.[CrossRef][Medline] [Order article via Infotrieve] Humphries JD, Byron A, Humphries MJ. Integrin ligands at a glance. J Cell Sci. 2006;119:3901–3903.[Free Full Text] Brooks PC, Clark RA, Cheresh DA. Requirement of vascular integrin alpha v beta 3 for angiogenesis. Science. 1994;264:569–571.[Abstract/Free Full Text] Friedlander M, Brooks PC, Shaffer RW, Kincaid CM, Varner J, Cheresh DA. Definition of two angiogenic pathways by distinct v integrins. Science. 1995;270:1500–1502.[Abstract/Free Full Text] Francis SE, Goh KL, Hodivala-Dilke K, et al. Central roles of alpha5beta1 integrin and fibronectin in vascular development in mouse embryos and embryoid bodies. Arterioscler Thromb Vasc Biol. 2002;22:927–933.[Abstract/Free Full Text] Pozzi A, Moberg PE, Miles LA, Wagner S, Soloway P, Gardner HA. Elevated matrix metalloprotease and angiostatin levels in integrin alpha 1 knockout mice cause reduced tumor vascularization. Proc Natl Acad Sci U S A. 2000;97:2202–2207.[Abstract/Free Full Text] Pozzi A, LeVine WF, Gardner HA. Low plasma levels of matrix metalloproteinase 9 permit increased tumor angiogenesis. Oncogene. 2002;21:272–281.[CrossRef][Medline] [Order article via Infotrieve] Zweers MC, Davidson JM, Pozzi A, et al. Integrin alpha2beta1 is required for regulation of murine wound angiogenesis but is dispensable for reepithelialization. J Invest Dermatol. 2007;127:467–478.[CrossRef][Medline] [Order article via Infotrieve] Alghisi GC, Ruegg C. Vascular integrins in tumor angiogenesis: mediators and therapeutic targets. Endothelium. 2006;13:113–135.[CrossRef][Medline] [Order article via Infotrieve] Reynolds LE, Wyder L, Lively JC, et al. Enhanced pathological angiogenesis in mice lacking beta3 integrin or beta3 and beta5 integrins. Nat Med. 2002;8:27–34.[CrossRef][Medline] [Order article via Infotrieve] Hodivala-Dilke KM, McHugh KP, Tsakiris DA, et al. Beta-3-integrin-deficient mice are a model for Glanzmann thromasthenia showing placental defects and reduced survival. J Clin Invest. 1999;103:229–238.[Medline] [Order article via Infotrieve] Senger DR, Claffey KP, Benes JE, Perruzzi CA, Sergiou AP, Detmar M. Angiogenesis promoted by vascular endothelial growth factor: regulation through alpha1beta1 and alpha2beta1 integrins. Proc Natl Acad Sci U S A. 1997;94:13612–13617.[Abstract/Free Full Text] Chen X, Su Y, Fingleton B, et al. Increased plasma MMP9 in integrin alpha1-null mice enhances lung metastasis of colon carcinoma cells. Int J Cancer. 2005;116:52–61.[CrossRef][Medline] [Order article via Infotrieve] Chen X, Su Y, Fingleton B, et al. An orthotopic model of lung cancer to analyze primary and metastatic NSCLC growth in integrin alpha1-null mice. Clin Exp Metastasis. 2005;22:185–193.[CrossRef][Medline] [Order article via Infotrieve] Zhang Z, Ramirez NE, Yankeelov TE, et al. 2β1 integrin expression in the tumor microenvironment enhances tumor angiogenesis in a tumor cell-specific manner. Blood. 2008;111:1980–1988.[Abstract/Free Full Text] Pozzi A, Wary KK, Giancotti FG, Gardner HA. Integrin alpha1beta1 mediates a unique collagen-dependent proliferation pathway in vivo. J Cell Biol. 1998;142:587–594.[Abstract/Free Full Text] Springer TA, Wang JH. The three-dimensional structure of integrins and their ligands, and conformational regulation of cell adhesion. Adv Protein Chem. 2004;68:29–63.[Medline] [Order article via Infotrieve] Partridge AW, Liu S, Kim S, Bowie JU, Ginsberg MH. Transmembrane domain helix packing stabilizes integrin alphaIIbbeta3 in the low affinity state. J Biol Chem. 2005;280:7294–7300.[Abstract/Free Full Text] Luo BH, Carman CV, Takagi J, Springer TA. Disrupting integrin transmembrane domain heterodimerization increases ligand-binding affinity, not valency or clustering. Proc Natl Acad Sci U S A. 2005;102:3679–3684.[Abstract/Free Full Text] Luo BH, Springer TA, Takagi J. A specific interface between integrin transmembrane helices and affinity for ligand. PLoS Biol. 2004;2:e153.[CrossRef][Medline] [Order article via Infotrieve] Schneider D, Engelman DM. Involvement of transmembrane domain interactions in signal transduction by alpha/beta integrins. J Biol Chem. 2004;279:9840–9846.[Abstract/Free Full Text] Lu CF, Springer TA. The alpha subunit cytoplasmic domain regulates the assembly and adhesiveness of integrin lymphocyte function-associated antigen-1. J Immunol. 1997;159:268–278.[Abstract] De Melker AA, Kramer D, Kuikman I, Sonnenberg A. The two phenylalanines in the GFFKR motif of the integrin alpha6A subunit are essential for heterodimerization. Biochem J. 1997;328:529–537.[Medline] [Order article via Infotrieve] Hughes PE, O'Toole TE, Ylanne J, Shattil SJ, Ginsberg MH. The conserved membrane-proximal region of an integrin cytoplasmic domain specifies ligand-binding affinity. J Biol Chem. 1995;270:12411–12417.[Abstract/Free Full Text] Vinogradova O, Velyvis A, Velyviene A, et al. A structural mechanism of integrin alpha(IIb)beta(3) "inside-out" activation as regulated by its cytoplasmic face. Cell. 2002;110:587–597.[CrossRef][Medline] [Order article via Infotrieve] Ulmer TS, Yaspan B, Ginsberg MH, Campbell ID. NMR analysis of structure and dynamics of the cytosolic tails of integrin alpha IIb beta 3 in aqueous solution. Biochemistry. 2001;40:7498–7508.[CrossRef][Medline] [Order article via Infotrieve] Wegener KL, Partridge AW, Han J, et al. Structural basis of integrin activation by talin. Cell. 2007;128:171–182.[CrossRef][Medline] [Order article via Infotrieve] Tadokoro S, Shattil SJ, Eto K, et al. Talin binding to integrin beta tails: a final common step in integrin activation. Science. 2003;302:103–106.[Abstract/Free Full Text] Kim M, Carman CV, Springer TA. Bidirectional transmembrane signaling by cytoplasmic domain separation in integrins. Science. 2003;301:1720–1725.[Abstract/Free Full Text] Moser M, Nieswandt B, Ussar S, Pozgajova M, Fassler R. Kindlin-3 is essential for integrin activation and platelet aggregation. Nat Med. 2008;14:325–330.[CrossRef][Medline] [Order article via Infotrieve] Ma YQ, Qin J, Wu C, Plow EF. Kindlin-2 (Mig-2): a co-activator of beta3 integrins. J Cell Biol. 2008;181:439–446.[Abstract/Free Full Text] Montanez E, Ussar S, Schifferer M, et al. Kindlin-2 controls bidirectional signaling of integrins. Genes Dev. 2008;22:1325–1330.[Abstract/Free Full Text] O'Toole TE, Mandelman D, Forsyth J, Shattil SJ, Plow EF, Ginsberg MH. Modulation of the affinity of integrin alpha IIb beta 3 (GPIIb-IIIa) by the cytoplasmic domain of alpha IIb. Science. 1991;254:845–847.[Abstract/Free Full Text] Hughes PE, Diaz-Gonzalez F, Leong L, et al. Breaking the integrin hinge: a defined structural constraint regulates integrin signaling. J Biol Chem. 1996;271:6571–6574.[Abstract/Free Full Text] Arias-Salgado EG, Lizano S, Shattil SJ, Ginsberg MH. Specification of the direction of adhesive signaling by the integrin beta cytoplasmic domain. J Biol Chem. 2005;280:29699–29707.[Abstract/Free Full Text] Mattila E, Pellinen T, Nevo J, Vuoriluoto K, Arjonen A, Ivaska J. Negative regulation of EGFR signalling through integrin-alpha1beta1-mediated activation of protein tyrosine phosphatase TCPTP. Nat Cell Biol. 2005;7:78–85.[CrossRef][Medline] [Order article via Infotrieve] Wary KK, Mainiero F, Isakoff SJ, Marcantonio EE, Giancotti FG. The adaptor protein Shc couples a class of integrins to the control of cell cycle progression. Cell. 1996;87:733–743.[CrossRef][Medline] [Order article via Infotrieve] Ginsberg MH, Partridge A, Shattil SJ. Integrin regulation. Curr Opin Cell Biol. 2005;17:509–516.[CrossRef][Medline] [Order article via Infotrieve] Loster K, Vossmeyer D, Hofmann W, Reutter W, Danker K. alpha1 Integrin cytoplasmic domain is involved in focal adhesion formation via association with intracellular proteins. Biochem J. 2001;356:233–240.[CrossRef][Medline] [Order article via Infotrieve] Vossmeyer D, Hofmann W, Loster K, Reutter W, Danker K. Phospholipase Cgamma binds alpha1beta1 integrin and modulates alpha1beta1 integrin-specific adhesion. J Biol Chem. 2002;277:4636–4643.[Abstract/Free Full Text] Peng L, Jin G, Wang L, Guo J, Meng L, Shou C. Identification of integrin alpha1 as an interacting protein of protein tyrosine phosphatase PRL-3. Biochem Biophys Res Commun. 2006;342:179–183.[CrossRef][Medline] [Order article via Infotrieve] Briesewitz R, Epstein MR, Marcantonio EE. Expression of native and truncated forms of the human integrin alpha 1 subunit. J Biol Chem. 1993;268:2989–2996.[Abstract/Free Full Text] Kinsella TM, Nolan GP. Episomal vectors rapidly and stably produce high-titer recombinant retrovirus. Hum Gene Ther. 1996;7:1405–1413.[Medline] [Order article via Infotrieve] Ireton RC, Davis MA, van Hengel J, et al. A novel role for p120 catenin in E-cadherin function. J Cell Biol. 2002;159:465–476.[Abstract/Free Full Text] Chen D, Roberts R, Pohl M, et al. Differential expression of collagen- and laminin-binding integrins mediates ureteric bud and inner medullary collecting duct cell tubulogenesis. Am J Physiol Renal Physiol. 2004;287:F602–F611.[Abstract/Free Full Text] Klekotka PA, Santoro SA, Zutter MM. alpha 2 integrin subunit cytoplasmic domain-dependent cellular migration requires p38 MAPK. J Biol Chem. 2001;276:9503–9511.[Abstract/Free Full Text] Klekotka PA, Santoro SA, Wang H, Zutter MM. Specific residues within the alpha 2 integrin subunit cytoplasmic domain regulate migration and cell cycle progression via distinct MAPK pathways. J Biol Chem. 2001;276:32353–32361.[Abstract/Free Full Text] Gardner H, Kreidberg J, Koteliansky V, Jaenisch R. Deletion of integrin alpha 1 by homologous recombination permits normal murine development but gives rise to a specific deficit in cell adhesion. Dev Biol. 1996;175:301–313.[CrossRef][Medline] [Order article via Infotrieve] Pozzi A, Macias-Perez I, Abair T, et al. Characterization of 5,6- and 8,9-epoxyeicosatrienoic acids (5,6- and 8,9-EET) as potent in vivo angiogenic lipids. J Biol Chem. 2005;280:27138–27146.[Abstract/Free Full Text] Smerling C, Tang K, Hofmann W, Danker K. Role of the alpha(1) integrin cytoplasmic tail in the formation of focal complexes, actin organization, and in the control of cell migration. Exp Cell Res. 2007;313:3153–3165.[CrossRef][Medline] [Order article via Infotrieve] O'Toole TE, Mandelman D, Forsyth J, Shattil SJ, Plow EF, Ginsberg MH. Modulation of the affinity of integrin IIbβ3 (GPIIb-IIIa) by the cytoplasmic domain of IIb. Science. 1991;254:845–847.[Abstract/Free Full Text] Czuchra A, Meyer H, Legate KR, Brakebusch C, Fassler R. Genetic analysis of beta1 integrin "activation motifs" in mice. J Cell Biol. 2006;174:889–899.[Abstract/Free Full Text] Luo BH, Carman CV, Springer TA. Structural basis of integrin regulation and signaling. Annu Rev Immunol. 2007;25:619–647.[CrossRef][Medline] [Order article via Infotrieve] Vossmeyer D, Kaufmann C, Loster K, et al. The cytoplasmic domain of the alpha1 integrin subunit influences stress fiber formation via the conserved GFFKR motif. Exp Cell Res. 2000;256:321–327.[CrossRef][Medline] [Order article via Infotrieve] McMullen ME, Bryant PW, Glembotski CC, Vincent PA, Pumiglia KM. Activation of p38 has opposing effects on the proliferation and migration of endothelial cells. J Biol Chem. 2005;280:20995–21003.[Abstract/Free Full Text] Lamalice L, Houle F, Huot J. Phosphorylation of Tyr1214 within VEGFR-2 triggers the recruitment of Nck and activation of Fyn leading to SAPK2/p38 activation and endothelial cell migration in response to VEGF. J Biol Chem. 2006;281:34009–34020.[Abstract/Free Full Text] Ivaska J, Reunanen H, Westermarck J, Koivisto L, Kahari VM, Heino J. Integrin alpha2beta1 mediates isoform-specific activation of p38 and upregulation of collagen gene transcription by a mechanism involving the alpha2 cytoplasmic tail. J Cell Biol. 1999;147:401–416.[Abstract/Free Full Text] Liu H, Radisky DC, Wang F, Bissell MJ. Polarity and proliferation are controlled by distinct signaling pathways downstream of PI3-kinase in breast epithelial tumor cells. J Cell Biol. 2004;164:603–612.[Abstract/Free Full Text] Bastian P, Posch B, Lang K, et al. Phosphatidylinositol 3-kinase in the G protein-coupled receptor-induced chemokinesis and chemotaxis of MDA-MB-468 breast carcinoma cells: a comparison with leukocytes. Mol Cancer Res. 2006;4:411–421.[Abstract/Free Full Text] Gliki G, Wheeler-Jones C, Zachary I. Vascular endothelial growth factor induces protein kinase C (PKC)-dependent Akt/PKB activation and phosphatidylinositol 3'-kinase-mediates PKC delta phosphorylation: role of PKC in angiogenesis. Cell Biol Int. 2002;26:751–759.[CrossRef][Medline] [Order article via Infotrieve] Maffucci T, Piccolo E, Cumashi A, et al. Inhibition of the phosphatidylinositol 3-kinase/Akt pathway by inositol pentakisphosphate results in anti-angiogenic and antitumor effects. Cancer Res. 2005;65:8339–8349.[Abstract/Free Full Text] CiteULike    Connotea    Del.icio.us    Digg    Reddit    Technorati    What's this? This Article Abstract Full Text (PDF) Supplemental Figures All Versions of this Article: blood-2007-12-126433v1 112/8/3242    most recent Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Rights and Permissions Citing Articles Citing Articles via CrossRef Citing Articles via Google Scholar Google Scholar Articles by Abair, T. D. Articles by Pozzi, A. Search for Related Content PubMed PubMed Citation Articles by Abair, T. D. Articles by Pozzi, A. 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