To test the idea that interactors besides KRIT1 might be important for HEG1 function, we used wild-type (WT) HEG1 cytoplasmic tail and HEG1 lacking the C-terminal Tyr-Phe (∆YF) required for KRIT1 binding (Gingras et al., 2012) as an affinity matrix (Figure 1A and Figure 1—figure supplement 1A). To discern potential endothelial-specific interactors, we analyzed the proteins bound to these matrices from lysates of Human Umbilical Vein Endothelial Cells (HUVEC) or HeLa cells by mass spectrometry (MS; Raw data is deposited in the MassIVE repository housed at the Center for Computational Mass Spectrometry at UCSD. See Materials and methods section for more information). This analysis identified a Rap1 effector, Rasip1, among the proteins isolated from endothelial cells but not from HeLa cells (Figure 1), a result consistent with the endothelial-specific expression of Rasip1 (Mitin et al., 2006; Mitin et al., 2004). In contrast KRIT1, and the known KRIT1 interactors CCM2 and ITGB1BP1 (ICAP1) (Zawistowski, 2005; Zawistowski, 2002) were isolated from both lysates. All of these interactions were specific in that these proteins were detected at 35–125 fold less abundance in a control affinity matrix generated from the αIIb integrin. These proteins were also statistically significant according to SAINTexpress, a method for scoring protein-protein interaction data from affinity purification MS experiments. Moreover, deletion of the last 2 residues of the HEG1 tail abolished binding of KRIT1 as expected, and of the KRIT1-binding proteins CCM2 and ITGB1P1. In sharp contrast, Rasip1 binding was not reduced by the absence of the YF motif (Figure 1B and Figure 1—figure supplement 1B). Immunoblotting confirmed that endogenous Rasip1 interacts with both the HEG1 cytoplasmic tail and HEG1 ∆YF (Figure 1C). To ask whether Rasip1 and HEG1 can interact in living cells, we transfected HUVEC with mCherry-HEG1 cytoplasmic tail, containing a mitochondrial-targeting sequence (Figure 1D), and analyzed endogenous Rasip1 distribution. Targeting of mito-mCherry-HEG1, but not mito-mCherry alone, to mitochondria recruited Rasip1, demonstrating the interaction of HEG1 and Rasip1 in cells (Figure 1E). In addition, Rasip1 co-immunoprecipitated with full-length HEG1 (Figure 1F). Consistent with the mass spectrometry results, immunoblotting confirmed that Rasip1 bound to HEG1 ∆YF (Figure 1B,C), suggesting that the HEG1-Rasip1 interaction does not require KRIT1 binding to HEG1. To further test the KRIT1-dependence of the HEG1-Rasip interaction, we silenced KRIT1 expression (Figure 2A,B; 70% reduction in expression was achieved) and found that it did not affect binding of GFP-Rasip1 to the HEG1 cytoplasmic tail affinity matrix (Figure 2A) or Rasip1 recruitment to mito-mCherry-HEG1 (Figure 2C). Conversely, Rasip1 did not interact with the C-terminal 19 residues of HEG1, which are sufficient to bind KRIT1 (Figure 1—figure supplement 1C). Thus, we report that Rasip1 interacts with HEG1 independently of KRIT1.

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Rasip1 redistributes from the endothelial cell cytoplasm to cell-cell junctions during the process of junction assembly (Wilson et al., 2013). Because HEG1 is found in these junctions (Kleaveland et al., 2009), we hypothesized that the HEG1-Rasip1 interaction might mediate Rasip1 recruitment. To test this idea, we induced junctional assembly by disrupting adherens junctions with EGTA, followed by recalcification in the presence of the Epac1-specific cAMP analog 8-pCPT-2’-O-Me-cAMP (hereafter called '007') to activate Rap1 (Enserink et al., 2002). As expected, during junction assembly, Rasip1 was recruited to VE-cadherin-positive cell-cell contacts (Figure 3A,B). In sharp contrast, when HEG1 was silenced (Figure 3C) Rasip1 recruitment to cell-cell contacts was prevented (Figure 3A,B). To exclude off-target effects, silencing HEG1 expression with a second shRNA (Figure 3—figure supplement 1A) or an siRNA specific for HEG1 (Figure 3—figure supplement 1B) also prevented Rasip1 recruitment to EC contacts. Previous studies have shown that loss of HEG1-KRIT1 interaction increases Rho-Kinase (ROCK) activity (Gingras et al., 2012), however, a ROCK inhibitor, H-1152, did not restore Rasip1 recruitment in HEG1-silenced EC (Figure 3—figure supplement 2A,B). Moreover, silencing of KRIT1 (Figure 2B) did not prevent Rasip1 recruitment to cell-cell contacts during junction formation (Figure 3D). Thus, HEG1 mediates Rasip1 recruitment to forming EC junctions.

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At cell-cell contacts, Rasip1, in conjunction with the Rap1 effector Radil, suppresses RhoA/ROCK signaling via the RhoA RhoGAP ARHGAP29 (Post et al., 2013; Xu et al., 2011; Post et al., 2015). As HEG1 interacts with Rasip1, we tested whether HEG1 may also be involved in regulating Rasip1-Radil-ARHGAP29 complex formation. As expected, Radil and ARHGAP29 co-immunoprecipitated with Rasip1 (Figure 3—figure supplement 3A,B). Silencing HEG1 expression (Figure 3—figure supplement 3C; 75% reduction in mRNA expression was achieved) did not affect the interaction between Rasip1 and Radil or ARHGAP29 (Figure 3—figure supplement 3A, B). Next we tested whether HEG1 could regulate the interaction between Rasip1 and Rap1. Rasip1 interacted with constitutively active Rap1-V12; the interaction was not affected by silencing HEG1 expression (Figure 3—figure supplement 3C,D). Similarly, addition of purified HEG1 cytoplasmic tail peptide (5 µM) did not alter the interaction between Rap1-V12 and Rasip1 (Figure 3—figure supplement 3D). These data suggest that HEG1 is required for Rasip1 translocation to cell-cell contacts; however, formation of a Rasip1-Rap1 or Rasip1-Radil-ARHGAP29 complex is not dependent on the presence of HEG1.

To analyze the functional importance of Rasip1 binding to HEG1, we mapped the HEG1 region that binds to Rasip1 using truncations and deletions of the HEG1 cytoplasmic tail (Figure 4A–C). Deletion of 9 amino acids in HEG1 (aa 1327-1335; TDVYYSPTS) prevented binding of Rasip1 to HEG1 in vitro (Figure 4C), Rasip1 recruitment to mito-mCherry-HEG1 (Figure 4D), and co-immunoprecipitation of Rasip1 with murine HEG1(∆1283-1291) (corresponding to aa 1327-1335 in human HEG1; Figure 4E). Furthermore, a 22-mer encompassing this region (HEG1 1318-1339), which lacked KRIT1 binding, was sufficient to interact with Rasip1 (Figure 4B,C). Thus, we mapped the Rasip1 binding site in HEG1 to a 9 residue peptide and show that deletion of this sequence blocks the capacity of HEG1 to bind to and to recruit Rasip1.

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Full-length Rasip1 contains an N-terminal poly-Proline region and Ras Association (RA) domain, a central Forkhead-associated (FHA) domain, and a C-terminal Dilute (DIL) domain (Figure 5A). We transfected HEK293T cells with FLAG-tagged Rasip1(1-265; poly-Pro+RA), (266-550; FHA), or (551-963; DIL), and measured binding to HEG1 tail affinity matrix. Rasip1(266-550), containing the FHA domain, was sufficient for binding to HEG1 (Figure 5B). Next, we tested whether this region is necessary for binding to HEG1. Deletion of this region in Rasip1(∆334-539) (Figure 5A) disrupted HEG1 binding (Figure 5C). Thus, the region of Rasip1 encompassing the FHA domain is both necessary and sufficient to bind to HEG1. Furthermore, the interaction of HEG1 and the FHA domain was direct because purified Rasip1(266-550) bound to both HEG1, HEG1 ∆YF, or HEG1(1318-1339) peptide affinity matrices (Figure 5D). Thus, HEG1 binds directly to the FHA domain of Rasip1 via a 9 amino acid (TDVYYSPTS) region of HEG1.

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We studied recruitment of Rasip1 to assembling cell-cell junctions in HUVEC to test the functional relevance of the HEG1-Rasip1 interaction. As described above, during junction assembly, Rasip1 was recruited to endothelial cell-cell contacts. Deletion of the HEG1-binding central domain of Rasip1 abolished recruitment of Rasip1(∆334-539) to these junctions (Figure 5E). Rasip1 is known to associate with cytoplasmic vesicles (Xu et al., 2011; Mitin et al., 2004). Remarkably, Rasip1(∆334-539) accumulated on cytoplasmic vesicles, which often appeared to concentrate near cell-cell junctions; however, in contrast to the full length protein, Rasip1(∆334-539) never incorporated into the junctions. Thus, the region of Rasip1 that mediates its physical interaction with HEG1 is required for Rasip1 junctional localization.

Rasip1 regulates β1 integrin activation (Xu et al., 2011). To investigate whether HEG1 is involved in regulating β1 integrin activation as well, we silenced HEG1 or Rasip1 expression in HUVEC and measured the binding of 9EG7 monoclonal antibody as a reporter of β1 integrin activation (Lenter et al., 1993). As expected, silencing Rasip1 expression in HUVEC decreased 9EG7 binding. In contrast, silencing HEG1 expression did not affect levels of activated β1 integrin (Figure 6A) suggesting that the effect of Rasip1 on β1 integrin activation is independent of HEG1. Previous studies show that Rasip1 mediates Rap1 inhibition of RhoA activity and of the RhoA effector Rho kinase (ROCK). As a result, silencing Rasip1 expression in endothelial cells can increase phosphorylation of a ROCK substrate, myosin light chain 2 (MLC) (Xu et al., 2011), resulting in increased actin stress fibers and reduced cortical actin (Post et al., 2013). To test whether HEG1 is also involved in suppressing ROCK signaling we silenced HEG1 or Rasip1 expression in HUVEC and analyzed MLC phosphorylation. Similar to silencing Rasip1, silencing HEG1 expression in HUVEC by siRNA or shRNA increased phosphorylation of MLC and formation of stress fibers indicating increased ROCK signaling (Figure 6B,C). As both HEG1 and Rasip1 silencing resulted in increased MLC phosphorylation, we tested whether the interaction between HEG1 and Rasip1 is important for the regulation of RhoA/ROCK signaling. Silencing HEG1 expression in HUVEC increased MLC phosphorylation and formation of actin stress fibers, which was rescued by expressing shRNA-resistant murine HEG1 (Figure 6D). However, expression of Rasip1-binding deficient murine HEG1(∆1283-1291) (corresponding to aa 1327-1335 in human HEG1) failed to rescue the increase in MLC phosphorylation and actin stress fiber formation (Figure 6D). Flow cytometry analysis showed that both wild-type mHEG1 and mHEG1(∆1283-1291) were equally expressed in HUVEC (Figure 6—figure supplement 1). Conversely, silencing expression of Rasip1 increased MLC phosphorylation by ~40% (Figure 7A,B). Expression of shRNA-resistant wild-type Rasip1 (Figure 7—figure supplement 1) rescued increased MLC phosphorylation (Figure 7A,B). In contrast, HEG1-binding deficient Rasip1(∆334-539) failed to rescue increased MLC phosphorylation (Figure 7A,B) Moreover, expression of Rasip1(∆334-539) induced increased MLC phosphorylation even in the absence of Rasip1 silencing, indicating that it could act in a dominant negative manner (Figure 7C). These data show that the interaction between Rasip1 and HEG1 is important for regulation of RhoA/ROCK signaling and the actin cytoskeleton in endothelial cells.

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Silencing of Rasip1 increases endothelial permeability (Post et al., 2013; Post et al., 2015). As expected, silencing Rasip1 increased endothelial permeability, and this was rescued by expression of shRNA-resistant wild-type (WT) Rasip1 (Figure 7D,E). In contrast, HEG1-binding deficient Rasip1(∆334-539) failed to reverse increased endothelial cell permeability (Figure 7D,E). Thus, deletion of the HEG1 binding domain of Rasip1 abolishes the capacity of Rasip1 to translocate to cell-cell contacts, to inhibit ROCK, and to maintain EC junctional integrity.

As noted above, Rasip1 binds Rap1 and activation of Rap1 facilitates Rasip1 targeting to cell junctions and suppresses ROCK activity. To test whether Rap1 binding to Rasip1 is required for Rasip1 function and to assess the role of Rap1 in regulating the interaction of Rasip1 with HEG1, we analyzed the Rasip1 Ras-association (RA) domain. Comparison of the Rasip1 RA domain sequence showed that it was similar to those found in RalGDS, Afadin6, and Radil. Sequence alignment of Rasip1 RA domain with Radil shows a highly conserved sequence (31% identical, Figure 8A). A preliminary homology model of the Rasip1 RA domain closely resembles the structure of Radil RA domain (PDB ID 3EC8) (backbone rmsd 1.02Å for 126 a.a., Figure 8B). The Radil crystal structure shows a dimer that is stabilized by an extensive interface burying 30% (2,324 Ų) of the total surface area of each monomer (Figure 8B, highlighted in blue). Analysis of our model shows that the dimer interface is conserved in Rasip1 (Figure 8B, magenta and yellow), suggesting that it could also form a dimer similar to Radil RA domain and therefore could bind two Rap1 molecules per dimer (Figure 8C). Indeed, purified recombinant Rasip1 RA domain (residues 134-285) eluted earlier than expected from its monomer mass (16.3kDa) in size exclusion chromatography (SEC), suggesting that it forms a dimer (~35kDa) in solution (Figure 8D; green). The purified Rasip1 RA domain bound activated Rap1B (Figure 8D; red) and Isothermal Titration Calorimetry (ITC) measured a k_d = 0.77 ± 0.09 μM for the interaction (Figure 8E).

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On the basis of the homology model, we created a Rasip1 RA domain mutant (R182E) mutant that could reduce Rap1 binding and tested its interaction with Rap1. SEC revealed that Rasip1(R182E) exhibited a markedly reduced affinity for Rap1 (Figure 8D; black). Moreover, SEC showed that the Rasip1(R182E) mutant was well folded and exhibited the same elution pattern in SEC as the wild-type protein. Thus, we conclude that Rasip1 RA domain (R182E) remains well folded as a multimer but binds Rap1 with drastically reduced affinity. We examined the capacity of Rasip1(R182E) to localize with HEG1 in cells and found that this mutant was not targeted to mito-mCherry-HEG1 (Figure 8F). Thus, in living cells, Rasip1 recruitment to HEG1 is lost when a point mutant disrupts its capacity to bind Rap1. Rasip1(R182E) also failed to rescue the effect of silencing of Rasip1 on increased MLC phosphorylation or endothelial permeability (Figure 9A–C) although it was well expressed (Figure 9D). Thus, Rasip1(R182E) is well expressed and folded, binds to activated Rap1 with much reduced affinity and loses the capacity to interact with HEG1 in living cells, and to inhibit ROCK and maintain EC junctional integrity.

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Cloning of point mutants, deletions, and truncations was done by using Infusion-HD Eco Dry Cloning Kit from Clontech (Mountain View, CA). Wild-type HEG1 cytoplasmic tail (1274-1381), 1364X, C19, and αIIb intracellular tail were previously described (Gingras et al., 2012). His6-tagged HEG1 cytoplasmic tail peptides C54, C49, 1334X, 1328X, 1318-1339, and ∆1327-1335 containing an in vivo biotinylation peptide tag at the N-terminus were cloned into pET15b. mCherry-HA-Radil was a kind gift from professor S. Angers (University of Toronto, Canada). pcDNA3.1-YFP-ARHGAP29 was a kind gift from professor J.L. Bos (University of Utrecht, The Netherlands). Using pcDNA3.1-YFP-Rasip1 (a kind gift from professor J.L. Bos, University of Utrecht, the Netherlands) as a template for PCR, Rasip1 full-length, 2-265, 266-550, and 551-963 were cloned into p3xFLAG-CMV7.1 or pEGFP-C1 using EcoRI and BamHI restriction sites. Rasip1 ∆334-539 and R182E were generated with two-fragment PCR Infusion cloning using wild-type Rasip1 as a template and cloned into p3xFLAG-CMV7.1 using EcoRI and BamHI restriction sites. Constitutively active Rap1-V12 was cloned into p3xFLAG-CMV7.1 or pEGFP-C1 using EcoRI and BamHI restriction sites Using pcDNA3.1-SNP-FLAG-mHEG1 FL-V5 (modified from a kind gift from professor M.L. Kahn, University of Pennsylvania) as a template, wild-type FLAG-tagged murine HEG1 and ∆1283-1291 (corresponding to aa 1327-1335 in human HEG1) were cloned into pLVX-Het-1 lentiviral expression vector using EcoRI and MluI restriction sites. mHEG1-expressing lentiviral particles were prepared by co-transfection of pLVX-Het-1 mHEG1 WT or ∆1283-1291 with pMDLg/pRRE, pRSV-Rev, and pMD2.G in HEK293T cells. Supernatant, containing lentiviral particles, was collected. Wild-type Rasip1, R182E, and ∆334-539, were cloned into 2K7 lentiviral vector (A kind gift from Alexander Zambon, UC San Diego, USA) using BamHI and XhoI restriction sites. Rasip1-expressing lentiviral particles were prepared by co-transfection of 2K7-Rasip1 wild-type, R182E, or ∆334-539 with pMDLg/pRRE, pRSV-Rev, and pMD2.G in HEK293T cells. Supernatant, containing lentiviral particles, was collected. Mito-mCherry (Lenter et al., 1993) and mito-mCherry-HEG1 wild-type or ∆1327-1335 were cloned into pcDNA3.1(-) using XhoI and NheI restriction sites. In vitro transcription to generate mRNA was performed using mMESSAGE mMACHINE® T7 ULTRA Transcription Kit (AM1345, Life Technologies, Grand Island, NY) according to the manufacturer’s recommendations.

For lentiviral delivery, HUVEC were grown to 80% confluence on gelatin-coated glass coverslips, and then infected with 2K7-Rasip1-containing lentiviral particles. 72 Hours post-infection, HUVECs were prepared for immunofluorescence analysis (described below). For in vitro binding studies, Rasip1 266-550 was cloned into pETM-41 (EMBL) containing His6 tag and Maltose Binding Protein (MBP) tag using NcoI and BamHI restriction sites.

Oligo’s for shRasip1 (Clone TRCN0000437267, target sequence: CCACTGAGTTCTTCCGGAAAC), shHEG1#1 (Clone TRCN0000253693, target sequence: ACCTTCGTGACAGAGTTTAAA), and shHEG1#2 (Clone TRCN0000253694, target sequence: CATTGGGAGATAGGAGTTATT) are based on the public TRC (The RNAi Consortium, Broad Institute) library and cloned into pLKO.1 using EcoRI and Age1 restriction sites. shRNA-expressing lentiviral particles were prepared by co-transfection of pLKO.1-Rasip1, HEG1#, or HEG1#2 with pMDLg/pRRE, pRSV-Rev, and pMD2.G in HEK293T cells. Supernatant, containing lentiviral particles, was collected. For shRNA delivery, HUVEC were grown to 80% confluence on gelatin-coated glass coverslips, and then infected with shRNA-containing lentiviral particles. 72 Hours post-infection, HUVECs were prepared for immunofluorescence analysis (described below). siRNA specific for HEG1 was from Ambion (Silencer select, ID: s33148). HUVECs were grown to 80% confluence on gelatin-coated glass coverslips, and then transfected with siRNA using Lipofectamine RNAi Max reagent (Life Technologies) according to the manufacturer’s recommendations. 72 Hours post-transfection, HUVECs were prepared for immunofluorescence analysis (described below).

HEK293T cells were maintained in Dulbecco’s modified Eagle’s medium (Corning, Tewksbury, MA) supplemented with 10% FBS (Sigma-Aldrich, Saint Louis, MO), 1% nonessential amino acids, 1% L-Glutamine, and 1% penicillin and streptomycin (all obtained from Invitrogen, Grand Island, NY). Primary Human Umbilical Vein Endothelial Cells (HUVEC) were purchased from Lonza and cultured in EGM2 medium, supplemented with singlequots (Lonza, Walkersville, MD).

HEK293T cells were transfected using TransIT-LT1 transfection reagents according to the manufacturer’s recommendations (Mirus Bio, Madison, WI). 24 Hours post-transfection, cells were prepared for Western blot analysis (described below). HUVEC were transfected using TransMessenger Transfection Reagent (301525, QIAGEN, Valencia, CA) according to the manufacturer’s recommendations.

Polyclonal anti-Rasip1 (Abcam, Cambridge, MA) was used for immunoblotting at 1:1000 and for immunofluorescence at 1:500. Mouse monoclonal (15B2) and rabbit polyclonal anti-KRIT1 (6832) were developed using recombinant KRIT1 FERM domain as the antigen, as previously described (Glading et al., 2007). Anti-KRIT1 15B2 was used for immunoprecipitation. Anti-KRIT1 (6832) was used for immunoblotting at 1:1000. Polyclonal-anti-pMLC (Cell Signaling) was used for immunoblotting at 1:1000 and for immune immunofluorescence at 1:300. Monoclonal-anti-VECadherin (Cell Signaling, Danvers, MA) was used for immunofluorescence at 1:1000. Polyclonal or monoclonal anti-FLAG (both from Sigma) were used for immunoblotting at 1:5000 and for immunofluorescence at 1:1000. Monoclonal anti-β-actin (Sigma) was used for immunoblotting at 1:5000. Monoclonal anti-GFP (Clontech) was used for immunoblotting at 1:5000. Monoclonal anti-HA (clone 16B12), monoclonal anti-human CD29 (Clone TS2/16, β1 integrin), and APC-conjugated anti-FLAG (clone L5) were from Biolegend (San Diego, CA). Rat anti-Mouse CD29 (Clone 9EG7, β1 integrin) was purchased from BD Biosciences (5 µg/ml) (San Jose, CA). Secondary AlexaFluor-labelled antibodies for immunofluorescence were from Life Technologies. Secondary AlexaFluor-labelled antibodies for Western blot analysis were from Rockland (Boyerton, PA) or Life Technologies. 8-pCPT-2’-O-Me-cAmp ('007') was purchased from Biolog (Hayward, CA) and used at 100 µM for 20 minutes. Rho Kinase inhibitor H-1152 was purchased from EMD biochemical (Billerica, MA) and used at 3 µM for 30 minutes.

HEG1 and αIIb intracellular tail proteins were prepared as previously described (Bear et al., 2000). In brief, His6-tagged tail proteins, containing an in vivo biotinylation peptide tag at the N-terminus, were expressed and purified from Escherichia coli. To perform pull-down assay using cell lysates, Human Umbilical Vein Endothelial Cells (HUVEC) and HEK293T, transfected as indicated, were lysed in cold lysis buffer (50 mM Tris-HCl, pH 7.4, 100 mM NaCl, 10 mM MgCl₂, 1%NP40, and 10% glycerol) plus protease and phosphatase inhibitor cocktails (Roche, Indianapolis, IN). A total of 10 µg of immobilized bead-bound proteins was added to 350 µg of clarified lysates. Reactions were kept at 4°C for 4 hours while rotating. After washing the beads with lysis buffer, samples were separated by SDS-PAGE. Bound proteins were detected by immunoblotting.

Human Rasip1, cloned in the pETM-11 (His-tagged) and pETM-41 (His-MBP-tagged) vectors, were expressed in Escherichia coli BL21 Star (DE3) cultured in LB media. Recombinant His-tagged Rasip1 RA domain (residues 134-285), wild-type and R182E polypeptides were purified by nickel-affinity chromatography following standard procedures. The His-tag was removed by cleavage with tobacco etch virus protease overnight, and the protein was further purified by size exclusion chromatography using a Superdex-75. The protein concentration for RA domain monomers was assessed using the A280 extinction coefficient of 24,980 M⁻¹. Recombinant His-MBP-fusion Rasip1 central domain (residues 266-550) was purified by nickel-affinity chromatography following standard procedures and dialyzed into PBS. The protein concentration for the His-MBP-Rasip1 central domain was assessed using the A280 extinction coefficient of 26,930 M⁻¹. Human Rap1 isoform Rap1B (residues 1–167) cloned into pTAC vector in the Escherichia coli strain CK600K was a generous gift of professor Alfred Wittinghoefer (Max Planck Institute of Molecular Physiology, Germany). The expression, purification and nucleotide exchange for GMP-PNP was achieved as described previously (Gingras et al., 2013). His-MBP-Rasip1 central domain at 2.5 μM was incubated with HEG1 matrices in binding buffer (PBS containing 0.1% Triton X-100, 0.5 mM EDTA, 2 mM DTT, and 1 mM Maltose). The mixture was incubated at 6°C for 1 hour on a rotating platform and the beads washed three times with ice cold binding buffer before electroporation. Subsequently, samples were separated by SDS-PAGE and analyzed by Coomassie-blue staining.

Human Umbilical Vein Endothelial Cells (HUVEC) were collected in cold lysis buffer (50 mM Tris-HCl, pH 7.4, 100 mM NaCl, 10 mM MgCl₂, 1%NP40, and 10% glycerol) plus protease and phosphatase inhibitor cocktails (Roche). A total of 2 µg of monoclonal anti-KRIT1 (15B2) antibody was added to 350 µg of clarified lysates and incubated at 4°C overnight while rotating. Protein G-Sepharose (Invitrogen) was added to the reaction mixture and further incubated for 4 hours at 4°C. After three washes with cold lysis buffer, beads were mixed with sample buffer and proteins were separated by SDS-PAGE. Bound KRIT1 was detected by using polyclonal anti-KRIT1 (6832) antibody.

HEK293T or U2OS cells, transfected and infected as indicated, were collected in cold lysis buffer (50 mM Tris-HCl, pH 7.4, 100 mM NaCl, 10 mM MgCl₂, 1%NP40, and 10% glycerol) plus protease and phosphatase inhibitor cocktails (Roche) and subsequently, clarified lysate was incubated with anti-FLAG G1 Affinity Resin (Genscript, Piscataway, NJ; L00432-1) and incubated at 4°C for 3 hours while rotating. After three washes with cold lysis buffer, beads were mixed with sample buffer and proteins were separated by SDS-PAGE.

Human Umbilical Vein Endothelial Cells (HUVEC) were grown to 80% confluence on gelatin-coated glass coverslips, then infected or transfected as indicated. After 72 hours, cells were fixed with 3.7% formaldehyde in Phosphate-Buffered-Saline (PBS) for 10 minutes (RT) and subsequently permeabilized with 0.1% Triton X-100 in PBS for 5 minutes (RT). Coverslips were then blocked for 1 hour (37°C) using 10% normal goat serum in PBS. Immunostainings were performed with the indicated antibodies (60 min; RT). Imaging was performed with a Perkin Elmer UltraView Vox Spinning Disk Confocal using a 40x/NA-1.30 or a 60x/NA-1.42 oil objective.

For the calcium switch assay, HUVEC were grown to 80% confluence on gelatin-coated glass coverslips, then infected/transfected as indicated. 72 Hours after infection cells were washed with cold PBS and then incubated with DMEM (5% FBS; 4 mM EGTA) for 30 minutes at 37°C. Cells were then washed with cold PBS and incubated with DMEM (2 mM CaCl; 100 µM 8-pCPT-2’-O-Me-cAmp) for 20 minutes at 37°C. Next, cells were stained as described above. When indicated, to inhibit Rho Kinase activity cells received ROCK inhibitor H-1152 (3 µM) for 30 minutes (37°C) prior to calcium switch assay.

In vitro permeability assay was performed as described previously (Pfaff et al., 1998). In brief, membrane filter inserts (pore size 0.4 µm, diameter 12 mm, Corning) were coated with fibronectin (10 µg/ml) for 1 hour at 37°C and placed in a 12-wells plate. Subsequently, fibronectin solution was removed and 1.2x10⁵ Human Umbilical Vein Endothelial Cells (HUVEC) were seeded in each insert. When 80% confluent, cells were infected as indicated and grown for 72 hours. Lower chamber medium was replaced with transport buffer (phenol-free DMEM, 2%FBS). Medium in upper chamber was replaced with transport buffer containing 2 mg/ml 70kD-FITC-Dextran (Sigma). Inserts were then incubated at 37°C and transported sequentially to new plates containing transport buffer at 5 minute intervals for 30 min. FITC in lower chamber medium was measured for each timepoint as well as a 1:1000 dilution of the remaining solution in the upper chamber. Permeability coefficient was calculated for each condition and then normalized as fold change in permeability compared to control HUVEC.

Lysis Buffer (50 mM Tris-HCl pH 7.4, 100 mM NaCl, 10 mM MgCl₂, 1% NP-40, 10% glycerol, 1 mM PMSF and 1x Sigma protease inhibitors) was added at a 1:4 ratio (pellet weight:volume) to a frozen cell pellet (approximately 300mg) of either HeLa or HUVEC cells. The pellet was resuspended by pipetting up and down. 1 μl of benzonase was added per sample and the samples were sonicated for 30secs at amplitude 0.37 (10s pulse, 2s off). The samples were then nutated for 30min at 4°C and subsequently centrifuged at 18K G for 20 min at 4°C. The supernatant was then passed through a 20 μm filter and transferred to a fresh tube. 20 μl of pre-bound streptavidin beads (bound with either HEG1, HEG1ΔYF or ITGA2B) were added to the filtered lysates and incubated for 2 hours at 4°C with gentle agitation. The beads were pelleted by centrifugation (1000g for 1 min), and the supernatant discarded. The beads were washed 1x with 500 μl of cold Lysis Buffer and 2x with 500 μl of cold Wash Buffer (50 mM Hepes-NaOH pH 8.0, 150 mM NaCl, 2 mM EDTA, 0.1% NP40, 10% glycerol). The beads were then transferred to a new tube in 500 μl of 20 mM Tris-HCl, pH 8, and washed again in 500 μl of Tris-HCl. The beads were resuspended in 100 μl of 50 mM ammonium bicarbonate, pH 8.0, containing 500 ng of trypsin, and the mixture was incubated at 37°C with agitation overnight (~15 hours). In the morning, another 500 ng of trypsin was added (in 20 μL of 20 mM Tris-HCl, pH 8), and the sample incubated at 37°C for an additional 4 hours. Finally, the supernatant was collected, and the beads washed twice in 40 μl of water. These washes were pooled with the collected supernatant. Formic acid was added to 2% to stop the digestion. The sample was subsequently vacuum-centrifuged to dryness and resuspended in 5 μl of 5% formic acid for LC/MS-MS. LC-MS/MS was performed using an Eksigent nano Ultra HPLC and a ThermoFisher Orbitrap Velos mass spectrometer. Samples were loaded using an autosampler directly onto a home-made 75 μm ID x 10cm packed tip column filled with C18 particles (3 μm, Reprosil, Dr. Maish). Digested peptides were separated over a 90min acetonitrile gradient (2-–35% buffer B over 90 min, 35-80% buffer B over 7 min, hold buffer B at 80% 8 min, and 80-2% B in 2 min) at 200 nl/min. Buffer A was water with 0.1% formic acid; buffer B was 100% ACN with 0.1% formic acid. A data dependent mode was used to acquire an MS scan (60,000 resolution) followed by 10 MS/MS CID scans at low resolution. Peptides were dynamically excluded for 15 sec after being selected for MS/MS. MS/MS spectra were queried against the RefSeq human database (version 53) using Mascot version 2.3.02 (Matrix Science). Methionine residues were searched with a variable modification of +15.9949 Da, and asparagine and glutamine with a variable modification of + 0.984016. Peptides were queried using tryptic cleavage constraints with a maximum of two missed cleavages sites. The mass tolerances were 12 ppm for parent masses and 0.6 Da for fragment masses. Experiments were performed in duplicate and significant interactors were determined using SAINTexpress (Lopez-Ramirez et al., 2012). Data was exported to an R-based dotplot tool for visualization (Teo et al., 2014).

All RAW mass spectrometry data and downloadable identification and SAINTexpress results tables are deposited in the MassIVE repository housed at the Center for Computational Mass Spectrometry at UCSD (http://proteomics.ucsd.edu/ProteoSAFe/datasets.jsp). The dataset has been assigned the MassIVE ID MSV000079420 and is available for FTP download at: ftp://MSV000079420@massive.ucsd.edu. The dataset was assigned the ProteomeXchange Consortium (http://proteomecentral.proteomexchange.org) identifier PXD003328.

Size exclusion chromatography of recombinant Rasip1 RA domain with Rap1 was performed using a Superdex-75 (10/300) GL (GE Healthcare, Piscataway, NJ) column at room temperature. The proteins, 50 μM each, were mixed in a volume of 100 μl and incubated for 30 min at room temperature before loading onto the column, which was pre-equilibrated with and run with TBS containing 3 mM MgCl₂.

ITC data were collected using a VP-ITC microcalorimeter (MicroCal Ltd., Northampton, MA) at 25°C and analyzed by fitting to a single-site binding equation using MicroCal Origin Software. For Rap1B binding, 335 μM GMP-PNP-bound Rap1B was titrated from the syringe into the sample cell containing 30μM Rasip1 RA domain. All proteins were dialyzed into 20 mM Sodium Phosphate, 200 mM NaCl, 3 mM MgCl₂, pH 6.5 containing 0.1 mM GMP-PNP before performing the experiments.

We generated an homology model of the Rasip1 RA domain using the Protein Homology/analogY Recognition Engine (PHYRE V 2.0) (Antonio Vizcaíno et al., 2015). The model with the highest confidence score was based on the Radil RA domain structure (PDB 3EC8, 100% confidence, 51% sequence identity). PISA (Proteins, Interfaces, Structures and Assemblies) software suggested that the biological assembly of the Radil RA domain structure is a dimer. The Radil dimer model was generated using the symmetry related molecule from the crystal structure and the Rasip1 RA domain dimer generated using those coordinates. The KRIT1 FERM F1 subdomain in complex with Rap1 (PDB 4HDO) was superimposed with the Rasip1 RA domain (RMSD 1.84 Å2 for 74 amino acids) to generate the Rasip1 RA domain complex with Rap1.

HUVEC were harvested by trypsinization, washed twice with PBS, and subsequently fixed in 2% paraformaldehyde for 10 min at room temperature. After three washes with PBS, cells were transferred into PBS supplemented with 0.5% BSA and 0.5% (wt:vol) saponin) for 20 min. Cells were centrifuged and resuspended in this buffer, containing APC-conjugated anti-FLAG antibody (1:200). After incubation on ice (30 min), cells were washed three times in buffer and then resuspended in PBS for analysis on a BD FACSCalibur (BD Biosciences, San Diego, CA, USA) using CellQuest software (BD Biosciences).

Cells were harvested by trypsinization, washed twice with PBS, washed once with Tyrode’s buffer (125 mM NaCl, 12 mM NaHCO₃, 2.5 mM KCl, 1 mM MgCl₂ and CaCl₂, 5 mM HEPES and Glucose, pH 7.4), then resuspended in Tyrode’s buffer containing primary antibodies (9EG7, TS2/16, Mouse IgG, or Rat IgG; 5 µg/ml), and incubated on ice for 30 min. Subsequently, cells were washed three times in Tyrode’s buffer, and incubated in Tyrode’s buffer containing goat anti-mouse or rat antibodies coupled to Alexa Fluor 647 (1:300) for 30 min on ice. Cells were further incubated with propidium iodide (1 μg/ml), then washed three times in Tyrode’s buffer, and analyzed on a BD FACSCalibur (BD Biosciences, San Diego, CA, USA) using CellQuest software (BD Biosciences). Level of 9EG7 (active β1) and TS2/16 (total β1) binding was corrected for background by substracting isotype control values. Results show 9EG7 binding relative to total β1 expression.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Wilma McLaughlin for assistance in molecular biology and Dr AA Bobkov for help with ITC.

© 2016, de Kreuk et al.

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